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Alma Mater Studiorum – Università di Bologna
DOTTORATO DI RICERCA IN
Scienze ambientali: tutela e gestione delle risorse naturali
Ciclo XXVII
Settore Concorsuale di afferenza: 03/A1 - CHIMICA ANALITICA
Settore Scientifico disciplinare: CHIM/01 - CHIMICA ANALITICA
TITOLO TESI
Biochar characterization for its environmental and
agricultural utilization. Occurrence, distribution and fate of
labile organic carbon and polycyclic aromatic hydrocarbons
Presentata da:
Alessandro Girolamo Rombolà
Coordinatore Dottorato
Relatore
Prof. Enrico Dinelli
Prof. Daniele Fabbri
Esame finale anno 2015
1. Introduction
1.1. Biochar System
1.1.1. Biochar definition
1.1.2. Biochar regulation
1.1.3. Learning from history
1.2. Potential biomass for biochar production
1.3. Biochar production techniques
1.4. Chemical reactions behind the production of hydrochar and biochar
1.5. Biochar characteristics
1.6. Environmental impact of biochar
1.6.1
Biochar and climate change
1.6.2. Biochar and soil
1.6.3. Biochar and waste management
1.6.4. Biochar and other environmental effects
1.7. Stability of biochar
1.8. Biochar and pollutants
1.9. Biochar: some non-negligible issues
2. Aim of the thesis
3. Determination of PAHs: method development and application
3.1. Determination of polycyclic aromatic hydrocarbons in biochar and biochar
amended soil
3.2. Quantitative determination of PAHs in an agricultural soil treated with biochar
4. Biochar characterization for agricultural utilization
4.1. Relationships between chemical characteristics and phytotoxicity of biochar from
poultry litter pyrolysis
5. Application of analytical pyrolysis methods to the characterization of organic
carbon in biochar
5.1. Characterisation of soil and biochar amended soil by hydropyrolysis
5.2. Characterization of biochar stability by hydropyrolysis and pyrolysis-GC/MS
6. Conclusions
References
Acknowledgments
1.
Introduction
1.1.
Biochar System
In the last half-century (1965-2015), the number of people on the planet has soared
from 3 billion to 7 billion (United Nations Population Division), placing ever-growing
pressure on the Earth and its resources.
Leading demographers, including those at the United Nations (UN) and the U.S.
Census Bureau, are projecting that world population will peak at 9.5 billion to 10 billion
later this century and then gradually decline as poorer countries develop. But what could
happen if those projections would be too optimistic? What if population continues to soar,
as it has in recent decades, and the world becomes home to 12 billion or even 16 billion
people by 2100, as a high-end UN estimate has projected? Such an outcome would clearly
have enormous social and environmental implications, including placing enormous stress
on the world’s food and water resources, spurring further loss of wild lands and
biodiversity, and hastening the degradation of the natural systems that support life on Earth
(Haub and Gribble, 2011). Other consequence of the rising world total population is a
tremendous demand and consumption of fossil fuels for energy generation and
consequently an increase in human-induced Greenhouse Gases (GHGs) emissions. The
world‫׳‬s total energy consumption was estimated at about 524 exajoules per year (EJ/y) and
has been predicted to increase by about 27% by the year 2020 and by about 65% by 2040
(BP statistical review, 2013) and (International energy outlook, 2014). The increase in
cost, depletion in availability, and deleterious environmental concerns associated with the
use of fossil fuels are the main topic of debates in energy meetings.
The urgency to address these threats creates an ever increasing demand for solutions
that can be implemented now or at least in the near future. These solutions need to be
widely implemented both locally by individuals and through large programmes in order to
produce effects on a global scale. This is a daunting and urgent task that cannot be
achieved by any single technology, but requires many different and integrated approaches
(Lehmann and Joseph, 2009).
Among the available options for these issues there is exploitation of chemical energy
captured into biomass by thermochemical conversions into energy, fuels and bioproducts.
Biomass can be converted to biofuels and bioproducts via thermochemical processes,
such as pyrolysis and gasification. The net carbon dioxide emissions from biofuel use are
considered virtually zero or negative because there leased CO2 was recycled from the
3
atmosphere captured during photosynthesis (Routa et al., 2012). In addition, since biomass
contains a low amount of sulphur and nitrogen, combustion of biofuels leads to lower
emissions of harmful gas, such as nitrous oxides (NOx) and sulphur dioxide (SO2), than
most of fossil fuels (Vassilev et al., 2010). Such advantages of biomass make it a
promising renewable energy resource.
The major products from biomass pyrolysis are a gaseous fraction (syngas), a liquid
material (bio-oil) and a solid residue (biochar) with yields that depend on the process
conditions. Syngas and bio-oil are considered as major intermediate products that can be
used to create fuels alternative to conventional fuels. Numerous studies have been
conducted involving up grading and utilization of syngas and bio-oil for various
applications (Noordermeer and Petrus, 2006; Kumar et al., 2010; Mortensen et al., 2011;
Swain et al., 2011).
Recently, biochar has received increasing attention for use in several applications.
Biochar has unique properties that make it a valuable soil amendment to sustainably
increase soil health and productivity, and also an appropriate tool for sequestering
atmospheric carbon dioxide in soils for the long term in an attempt to mitigate global
warming (Lehmann and Joseph, 2009). The recent broad interest in biochar has been
chiefly stimulated by the discovery that biochar is the primary reason for the sustainable
and highly fertile dark earths in the Amazon Basin, Terra Preta de Indio. Even though
biochar has been used in many other places at other times, and has even been the subject of
scientific investigation for at least a century, efforts have been isolated or regionally
focused (Lehmann and Joseph, 2009).
1.1.1. Biochar definition
According to Lehmann and Joseph (2009), biochar is defined as “a carbon (C)-rich
product when biomass such as wood, manure or leaves is heated in a closed container with
little or unavailable air” (Lehmann and Joseph, 2009). Shackley et al. (2012) defined
biochar more descriptively as “the porous carbonaceous solid produced by the
thermochemical conversion of organic materials in an oxygen depleted atmosphere that has
physicochemical properties suitable for safe and long-term storage of carbon in the
environment”. Verheijen et al. (2010) also defined biochar as “biomass that has been
pyrolyzed in a zero or low oxygen environment applied to soil at a specific site that is
expected to sustainably sequester C and concurrently improve soil functions under current
4
and future management, while avoiding short- and long-term detrimental effects to the
wider environment as well as human and animal health’’. The International Biochar
Initiative (IBI) standardized its definition as ‘‘a solid material obtained from the
thermochemical conversion of biomass in an oxygen-limited environment’’ (IBI, 2012).
While the European Biochar Certificate (EBC, 2014) defined biochar as “a heterogeneous
substance rich in aromatic carbon and minerals. It is produced by pyrolysis of sustainably
obtained biomass under controlled conditions with clean technology and is used for any
purpose that does not involve its rapid mineralisation to CO2 and may eventually become a
soil amendment”.
All of these definitions are directly or indirectly related to the biochar production
condition and its application to soil. Lehmann and Joseph (2009) distinguished biochar
operationally from charcoal. Primarily, the difference between these two terms lies in the
end use. The charcoal is a source of charred organic matter for producing fuel and energy
whereas the biochar can be applied for carbon sequestration and environmental
management. The term hydrochar is closely related to biochar; however, it is distinguished
by different condition like the hydrothermal carbonization of biomass (Libra et al., 2011).
In general, biochar is produced by dry carbonization or pyrolysis and gasification of
biomass, whereas hydrochar is produced as slurry in water by hydrothermal carbonization
of biomass under pressure. The two chars differ widely in chemical and physical properties
(Bargmann et al., 2013).
1.1.2. Biochar regulation
In 1984, Japan became the first country worldwide to approve the use of biochar as a
soil conditioner. For the first time in Europe, the Swiss Federal Ministry of Agriculture
officially approved the use of certified biochar in agriculture in 2013 (http://www.ithakajournal.net/schweiz-bewilligt-pflanzenkohle-zur-bodenverbesserung?lang=en). Approval is
based on strict, scientifically checked requirements with regard to the sustainability of
biochar production, to biochar quality and to user protection in its application. In the EU,
the use of biochar in agriculture is neither clearly regulated nor explicitly forbidden. In
Germany for example, the use of biochar as animal feed is allowed. It can thus be
composted with the manure and applied to fields. In addition, charcoal is allowed as an
additive for fertilizers and soil conditioners. What however is missing is an exact definition
5
of what can be counted as biochar and which production conditions and thresholds need to
be complied with. With the Swiss approval, we now have an exact definition, along with a
requirement for strict quality controls. Further, the European Biochar Certificate has been
developed to become the voluntary European industrial standard ensuring a sustainable
biochar production and low hazard use in agronomic systems (http://www.europeanbiochar.org/en).
In the United States (U.S.) some biochar production systems have been recommended
for generating C offsets by soil sequestration (De Gryze et al., 2010). Also, U.S. proposed
federal legislation to comprehensively address energy and climate change (i.e., the
American Power Act) included “projects for biochar production and use” to be considered
for domestic C offset programs (Gurwick et al., 2012). Recently, IBI certified the first
biochar material for effective use as a soil amendment for the development of small-scale
bio-refineries for the conversion of non-food biomass into biofuels and biochar in
California (http://www.biochar-international.org/certification). Biochar is commercially
available in the U.S. (Biochar Supreme, www.biocharsupreme.com; Biochar Solutions,
www.biocharsolutions.com). Further, large-scale biochar production from crop straw is
now commercially available in China (Pan et al., 2011). However, the biochar price is
claimed to be too much high (about 3.7 $ kg-1) and would not be balanced by the potential
economic gains based on average yield improvements and current prices for CO2 (Liu et
al., 2013). Thus, biochar has not yet made a substantial entry into large-scale agricultural
operations (IBI, 2014).
1.1.3. Learning from history
Several thousand years ago, pre-Columbian indigenous farmers used ‘slash and char’ to
bring soils into production. ‘Slash and char’ sequesters approximately 50% of the carbon
in the vegetation whereas ‘slash and burn’, still practiced by some cultures today,
sequesters only about 3% of the carbon. To create ‘char’, vegetation cleared from new
areas was smouldered at moderate temperatures in the absence of oxygen. The result was
then dug into the soil. Food scraps and waste materials were also added with the result that
the terra preta soils have not only high carbon (and are black) but also high fertility in
comparison with adjacent char-free soils (Fig. 1).
The terra preta soils are thought to have formed over a relatively short time span –
only 40-50 years. They range in depth from half a metre to two metres deep, and can
6
contain as much as 250 tonnes of carbon per hectare in the first 30 cm and 500 tonnes per
hectare up to one meter (but different authors have different values, indicating the
difficulty of assessing soil carbon, as explained in the Grassland newsletter article by
Parsons and Rowarth 2009). Unimproved soils from similar parent material have
approximately 60% less carbon than the ‘char-enriched’ soils.
Although stable, biochar is not inert – it can hold plant nutrients, including nitrogen,
and often has useful supplies of potassium, sulphur and phosphate in the accompanying
ash. It is probably this capacity that has resulted in reports that the addition of biochar
resulted in a doubling of crop production in South America. The biochar was added with
manure and food waste, as well as the ash resulting from the charring process. The ash also
has a liming effect, increasing soil pH. As the ash, manure and food waste was broken
down by microorganisms, the nutrients released that were not immediately immobilised
(by micro-organisms) or taken up by plants, were retained by the biochar instead of being
lost by leaching or, in the case of nitrogen, denitrification. Thus the biochar provided a
source of nutrients that did not come from the char, but were plant available?
Figure 1. Comparison of profiles of terra preta and adjacent soils (Source: IBI website).
7
1.2. Potential biomass for biochar production
The biomass potentials that could be available for biochar is categorized into two types:
(i) primarily produced biomass as a resource of bioenergy and biochar, and (ii) byproducts
as waste biomass. However, biochar production from dedicated crops could create
competition for land with any other land use option — such as food production or leaving
the land in its pristine state. Therefore, biochar should be made from biomass waste
materials. Appropriate biomass waste materials for biochar production include crop
residues (both field residues and processing residues such as nut shells, fruit pits, bagasse,
etc), as well as yard, food and forestry wastes and animal manures. Large amounts of
agricultural, municipal and forestry biomass are currently burned or left to decompose and
release CO2 and methane back into the atmosphere. They also can pollute local ground and
surface waters — a large issue for livestock wastes, therefore using these materials to
produce biochar allows to remove them from a pollution cycle.
In theory, any C-based feedstock can be pyrolysed to produce biochar, and so biochar
production has the potential to mitigate the increasing global problem of waste disposal.
To date, a wide range of waste streams have been considered and tested, including
biosolids (Chan and Xu, 2009), tannery wastes (Muralidhara, 1982), paper sludge
(Rajkovich et al., 2011) and sewage and wastewater sludge (Bridle and Pritchard 2004;
Hossain et al., 2010). The type of feedstock affects the properties of the resulting biochar
(Kloss et al., 2012) in terms of crop yield effects (Jeffery et al., 2011) and recalcitrance in
the soil (Zimmerman, 2010; Singh et al., 2012). Furthermore, it is likely to affect whether
the resulting biochar is classified as a waste product, with implications regarding its
permissibility for soil application (Sohi et al., 2010). Legislative issues surrounding
biochar application to soils produced from waste products, and the classification of such
biochar in terms of policy, is vital before its large-scale application can be implemented.
One readily apparent trade-off regarding choice of feedstock for biochar production is
the issue of stability of the resulting biochar vs. its nutrient content. For example, evidence
suggests that biochars prepared from poultry litter support greater increases in crop
productivity than those obtained from wood (Jeffery et al., 2011), probably because of a
higher nutrient contents in this feedstock. However, biochars from poultry litter are less
stable in the soil than those prepared from wood (Singh et al., 2012).
8
1.3. Biochar production techniques
The conversion of biomass into biochar can be performed with the help of a variety of
thermochemical
processes,
including
pyrolysis,
gasification
and
hydrothermal
carbonization (HTC). The choice of treatment method depends on type of feedstock (wet
or dry) and the desired properties of biochar for its different applications. In fact, the
properties of a given biochar strongly depend on the characteristics of each process and
also on the material to which the process is applied. Under all thermal treatments, biochar
is generally produced by heating biomass at high temperature in the absence or limited
supply of oxygen. Thermal treatments are classified based on their operating conditions
such as: severity of process parameters (mainly reaction time and temperature), pre- and
post-processing requirements like shaping, sizing, drying, cooling, condensation, etc.
(Mosier et al., 2005; Goyal et al., 2008; Manyà et al., 2012)
Pyrolysis
The most common method to produce biochar is pyrolysis. Pyrolysis is a
thermochemical decomposition process during which biomass is heated at elevated
temperature (300-650 °C) in the absence of oxygen. At these temperatures, organic
materials thermally decompose releasing a vapor phase and biochar. By cooling the
pyrolysis vapor, polar and high-molecular-weight compounds condense out as bio-oil
while low-molecular-weight volatile compounds, like CO, CO2, CH4 and H2 (Brownsort,
2009; Mohan et al., 2006), remain as syngas. Biochar generally has a high carbon content,
up to a half of the total carbon of the original organic matter. Bio-oil is generally a
hydrophilic liquid, containing many oxygenated compounds, and can be obtained as a
single aqueous phase or phase-separated (Demirbas and Arin, 2002). Syngas is generally
composed by carbon dioxide, carbon monoxide, methane, hydrogen and C2 hydrocarbons
in varying proportions.
Depending upon the reaction time, temperature, and heating rate the pyrolysis process
is sub-divided in four categories: slow, fast, flash and intermediate pyrolysis (Bridgwater
and Peacocke, 2000; Onay et al., 2003; Laird et al., 2009; Jones et al., 2009; Vamvuka,
2011).
Slow pyrolysis. Conventional or slow pyrolysis processes produces biochar by heating
biomass at a low heating rate for a relatively long residence time (Table 1) and usually at
lower temperature than fast pyrolysis (400°C). The target product is often the char, but this
9
is always accompanied by liquid and gas products although these are not always recovered.
Slow pyrolysis can be divided into traditional charcoal making and more modern
processes. In fact, this process has been practiced for thousands of years (Zhang et al.,
2010). It continues to be widely used for production of high quality charcoal for
metallurgical applications such as in the production of high grade silicon, as a leisure fuel
in many developed countries, and in developing countries as an essential and storable
commodity for cooking. There is widespread small scale local production in many
developed countries as a cottage industry but there are limited truly commercial operations
(a notable exception is charcoal production in Brazil for iron and steel production).
Several variables and factors play a critical role during the pyrolysis process, and
specifically: peak temperature, pressure, vapor residence time and moisture content (Antal
et al., 2003). The peak temperature is the highest temperature reached during the process.
As a general rule, the charcoal yield decreases as temperature increases. However, an
increase of the peak temperature results in an increase of the fixed-carbon content in
biochar (Schenkel et al., 1998; Antal et al., 2000; Antal et al., 2003a). This increase is
especially pronounced in the temperature range from 300 to 500 °C. In addition, the peak
temperature has influence on surface area and pore size distribution (both properties
generally related to specific adsorptive properties) of charcoals.
Fast pyrolysis. Fast pyrolysis produces biochar at a high heating rate (10-1000 °C s-1)
and short residence time (less than 10 s). The peak temperature is usually set between 500
and 550 °C in order to obtain the highest bio-oil yield (Maschio et al., 1999; Onay et al.,
2001; Yanik et al., 2007; Uzun et al., 2007). In this kind of pyrolysis, biomass decomposes
very quickly favouring the formation of bio-oil and inhibiting the formation of biochar
(about 15% of products) (Table 1).
Intermediate pyrolysis.
Intermediate pyrolysis operates between the reaction
conditions of slow and fast pyrolysis, including moderate heating rates up to 200-300 °C
min-1 and residence times for feedstock of 0.5-25 min. The product distribution generated
by this process is typically 40-60% of bio-oil, 20-30% syngas and 15-25% biochar. In
particular, the biochar obtained by intermediate pyrolysis is dry and has a brittle texture as
it contains less tar and therefore less toxic compounds making it suitable for further
applications, such as a solid fuel or as a soil amendment and/or as a fertilizer.
Flash pyrolysis. Flash pyrolysis occurs with very fast heating rates of ≥ 1000 °C s -1
and uses even shorter solid residence time (<0.5 s) than fast pyrolysis. The flash
carbonization process has been developed by Antal and Grönli (2003) at the University of
10
Hawaii as an efficient way to produce biochar by the ignition of flash fire at elevated
pressure in a packed bed of biomass. Air is used to pressurize a vessel to an initial pressure
of 1-2 MPa, and a flash fire is ignited at the bottom of a packed bed. After a few minutes,
air is delivered to the top of the packed bed and biomass is converted to charcoal. The total
reaction time is less than 30 min and the temperature profile of the packed bed is
conditioned by several factors: biomass feedstock, moisture content of the feedstock,
heating time and the total amount of air delivered (Antal et al., 2003b). In any case, the
flame front moves up the packed bed, causing the middle and top temperatures to
successively increase, until reaching values near 600 °C. This procedure determines a
significant improvement in yields with respect to conventional carbonization or slow
pyrolysis (Antal et al., 2003b; Nunoura et al., 2006).
Gasification
Gasification is an alternative thermo-chemical conversion technology suitable for
treatment of biomass or other organic matter including municipal solid wastes or
hydrocarbons such as coal. Gasification primarily transforms biomass into a gaseous
mixture (syngas containing CO, H2, CO2, CH4, and smaller quantities of higher
hydrocarbons) by supplying a controlled amount of oxidizing agent under high temperature
(> than 700 °C). Although they are designed to produce gas, gasifiers under some
conditions can also produce reasonable yields of char. Therefore they have been proposed
as an alternative production route to pyrolysis for biochar (Brown, 2009). The typical
biochar yield of gasification averages about 10wt% of biomass (Meyer et al., 2011; Qian et
al., 2013). The oxidizing agent used in gasification can be oxygen, air, steam or mixtures
of these gases. Air gasification produces syngas with low heating values of 4-7 MJ/Nm3,
while gasification with steam produces syngas with high heating values of 10-14 MJ/Nm3
(Kumar et al., 2009).
Hydrothermal carbonization
Hydrothermal carbonization (HTC) of biomass takes place in water at elevated
temperatures (160-800 °C). Since the water temperature is above 100 °C, the reaction
pressure also must be elevated (more than 1 atm) to maintain the water in a liquid form.
According to the reaction temperature, hydrothermal carbonization can be divided into
high-temperature HTC (between 300 and 800 °C) and low-temperature HTC (below 300
°C) (Hu et al., 2010). Since the reaction conditions of high-temperature HTC (above 300
11
°C) are beyond the stability condition of most organic compounds, the dominant reaction
in this case is hydrothermal gasification and the dominant products are gases, such as CH4
and H2 (Kruse et al., 2013). Below 300 °C, gasification is limited and carbonization of
biomass to char dominates the reaction. Low-temperature HTC can mimic the natural
coalification of biomass, although the reaction rate is higher and reaction time is shorter if
compared to the thousand/billion years of slow natural coalification of biomass. Char yield
of low-temperature biomass HTC varies from 30% to 60% depending on the feedstock
properties, reaction temperature and pressure. Since HTC requires water, this may be a
cost effective biochar production method for feedstock with high moisture content (Titirici
et al., 2012).
Table 1.
Temp &
Duration
Solid
(Biochar)
Liquid
(Bio oil)
Gas
(Syn Gas)
Slow Pyrolysis
~ 500C
min to days
35%
30%
35%
Intermediate
450-500°C
min
20%
50%
30%
Fast Pyrolysis
~ 500C
seconds
12%
75%
13%
Flash Pyrolysis
> 800°C
seconds
10%
75%
15%
Gasification
> 800C
hours
10%
5%
85%
180-250°C
1-12 hours
70%
25%
5%
HTC
1.4. Chemical reactions behind the production of hydrochar and biochar
During the production of biochar, biomass undergoes to a series of chemical reactions
that are highly complicated and depend on both the nature of the biomass and the
conditions (Glaser et al., 2001; Di Blasi, 2008; Babu, 2008; Funke and Ziegler, 2010).
However, most of these chemical reactions have similar thermochemical pathways, i.e. the
12
degradation and depolymerization of polymeric composition of biomass take place,
resulting in the formation of solid, liquid and gaseous (by-) products. The fundamental
difference in various thermochemical treatments lies in the operating conditions and
reaction medium that are used for the production of biochar and hydrochar. The highest
reaction temperature reached during a thermochemical process is the main parameter that
controls: i) the dominant reactions; ii) the reaction mechanism; iii) the physicochemical
properties of char. Decarboxylation, dehydration, de-carbonylation, de-methoxylation,
intermolecular rearrangement, condensation, aromatization, etc. are some of the proposed
chemical reactions that can take place (Funke et al., 2010). However, in real practice, it is
difficult to maintain uniform temperature profiles in pyrolysis reactors; therefore, it is most
likely possible that many of the aforementioned reaction mechanisms take place
simultaneously (Glaser et al., 2001).
The thermal stability of the polymeric constituent of lignocellulosic of biomass
significantly depends on the reaction medium in which the process is carried out. Under
standard pressure conditions (e.g. pyrolysis) the decomposition of hemicellulose takes
place between 200-300 °C, followed by cellulose that decomposes at higher temperatures
(300-400 °C). Lignin is the most thermo-chemically stable polymer and decomposes in a
wide temperature range peaking around 600 °C (Grønliet al., 2002) In contrast, during
HTC the degradation/depolymerization of biomass occurs at significantly lower
temperatures than pyrolysis (Yan et al., 2009). The degradation of hemicellulose and
cellulose under HTC process starts at around 160-180 °C, where most of the lignin still
remains stable until near or above critical point of water (Bobleter, 1994). The polymeric
degradation of biomass in the HTC process is controlled by reaction mechanisms very
similar to those in the pyrolysis process. However, due to the presence of hot compressed
water process the degradation of biomass during HTC is primarily initiated by hydrolysis,
resulting in the cleavage of ether and ester bonds between monomeric sugars by the
addition of one molecule of water (Bobleter, 1994) and thereby reducing the activation
energy levels of biomass polymers (Glaser et al., 2001). During HTC, the cellulose and
hemicellulose are partially or fully driven off, leaving behind a char with high lignin
content.
13
1.5. Biochar characteristics
The composition and the chemistry of biochar can be very different according to the
variety of feedstock that have been thermally degraded under a range of conditions (Antal
et al. 2000; Antal and Grønli 2003; Amonette and Joseph 2009; Krull et al. 2009; Libra et
al. 2011; Cantrell et al. 2012).
Kuwagaki (1990) proposed that seven properties should be measured for a quality
assessment for agronomically-used biochar: pH, volatile matter, ash content, water holding
capacity, bulk density, pore volume, and specific surface area. IBI and the EBC have
developed a series of guidelines for biochar production and quality (IBI, 2013; EBC,
2014). For instance, IBI sets a range of 6-20 mg kg-1dry weight (dw) as the maximum allowable
threshold values (varying between different countries) for the sum of the 16 US
Environmental Protection Agency’s (EPA) polycyclic aromatic hydrocarbons (PAHs) in
biochar (IBI, 2013). Similarly, EBC requires PAHs to be below 4 and 12 mg kg-1dw, in
premium and basic grade biochars, respectively (EBC, 2014). Both documents also list
guide values for a number of heavy metals, elemental contents (C, H, N, O) and their
molar ratios, and specific surface area (SSA) (Table A1 in supplementary materials). With
regard to carbon content, for instance, EBC proposed that the biochar's carbon content
must be higher than 50%dw. While IBI requires a carbon content higher than 60% and
30%dw for first and second class biochar, respectively.
The organic carbon content of pyrolysed chars ranges between ±5% and 95% of the dry
mass, dependent on the feedstock and process temperature used. Generally, biochars
derived from solid biomass residues tend to have higher carbon contents (63-82%) than
those derived from digestable biomass residues (35-66%) and digestates (42%) (Enders et
al., 2012). For instance the carbon content of pyrolysed poultry manure is around 35%
(Song et al., 2012), while that of wood is around 70-80% (Fabbri et al., 2012). When using
mineral-rich feedstocks such as sewage sludge or animal manure, the pyrolysed products
tend to have high ash content.
At low temperature, biochar chemical composition is closer to the original feedstock
while high temperature biochar is similar to graphite (Masiello, 2004). The biochars
produced at around 350 °C are mainly dominated by aromatic (aryl) carbon with small
proportions of alkyl-O and alkyl-C. When the reaction temperature is further increased
(>500 °C), these alkyl-O and alkyl-C were completely converted to aryl-C and these chars
usually have very low H/C ratios. In general, the carbon content of biochar is inversely
14
related to biochar yield. Increasing pyrolysis temperature from 300 to 800°C decreases the
yield of biochar from 67 to 26% and increases the carbon content from 56 to 93% (Tanaka,
1963). Beyond a certain threshold, the mass of biochar may decrease without any affect on
the amount of carbon retained within it; but as mass is lost, the ash content of biochar
increases (Bourke et al., 2007). Pyrolysis temperature greatly affects the surface area of
pyrolysis products. In particular, the increase of pyrolysis temperature determines an
increase of surface area of biochar. This effect of temperature suggest that biochar
prepared at low temperature may be suitable for controlling the release of fertiliser
nutrients (Day et al., 2005), whilst high temperature biochars would be more suitable as
activated carbon (Ogawa et al., 2006). The surfaces of low temperature biochar are,
however, hydrophobic and this may limit the capacity to store water in soil. The scanning
electron microscopy images (Fig. 2) of these biochars clearing show that have a structure
with voids and micropores in which water can be retained.
It is critically important to characterize biochar because its characterization will play a
vital role in determining its importance and application in the industry and environment.
For example, a biochar with low carbon content and high ash content is not suitable for
energy product, and in the same way a biochar with low surface area and low adsorption
capacity is not meant for agricultural and wastewater treatment applications.
Figure 2. Scanning electron microscopy images of orchard pruning biochar at
magnification × 500 and 7000.
15
1.6. Environmental impact of biochar
1.6.1. Biochar and climate change
The greenhouse-gas (GHG) concentrations of CO2, CH4 and NOx in the atmosphere
have strongly risen since pre-industrial times (Ciais et al., 2013). The driver of these
concentrations is an increase in human-induced GHG emissions (Ciais et al. 2013).
Emissions of CO2 are attributed to the increased use of fossil fuels, as well as to the
enhanced clearing and burning of forests (Fearnside, 2000), and the expanding of
agriculture. As the main cause for the rise in the global mean surface temperature (Bindoff
et al. 2013), it is widely recognized that the anthropogenic GHG concentrations need to be
drastically reduced to combat climate change. The world is on a trajectory that results in a
level of emissions consistent with long-term average temperature increase of more than 3.5
°C (International Energy Agency, 2011). To change this trajectory, a timely and ambitious
programme of mitigation measures is needed. Several studies have shown that, to stabilize
global mean surface temperature, cumulative anthropogenic GHG emissions must be kept
below a maximum upper limit, thus indicating that future net anthropogenic emissions
must approach zero.
16
Figure 3. Scheme of biochar driven soil carbon sequestration. Diagram from Nature
Publishing Group (Lehmann, 2007).
production of biochar, in combination with its storage in soils, has been suggested
as one possible
Biochar’s climate-mitigation potential primarily
stems from its highly recalcitrant nature (
), which
slows the rate at which photosynthetically fixed carbon (C) is returned to the atmosphere.
The biochar also improves soil fertility, stimulating plant growth, which then consumes
more CO2 in a feedback effect and the energy generated as part of biochar production can
displace carbon positive energy from fossil fuels.
Moreover, biochar applied to soils has been shown to reduce NOx emissions
significantly (with the added benefit of reducing nitrogen fertiliser requirements). As NOx
are approximately 320 times more effective as a GHG than CO2, biochar could be very
important in mitigating emissions. However, the mechanisms and quantities involved are
17
still being investigated.
Additional effects from adding biochar to soil can further reduce greenhouse gas
emissions and enhance carbon storage in soil. These include: (i) Biochar reduces the need
for fertilizer, resulting in reduced emissions from fertilizer production; (ii) Biochar
increases soil microbial life, resulting in more carbon storage in soil; (iii) Turning
agricultural waste into biochar reduces methane (another potent greenhouse gas) generated
by the natural decomposition of the waste. Moreover, there may be additional benefits
arising from the contribution of biochar to facilitate agricultural development and
improving the socioeconomic circumstances of farmers in developing countries.
Globally, Woolf et al. (2010) estimated that the potential impact of biochar for climatechange mitigation is 12% of current anthropogenic CO2-C equivalent (CO2-Ce) emissions
(that is, 1.8 Pg CO2-Ce per year of the 15.4 Pg CO2-Ce emitted annually), and that over the
course of a century, the total net offset from biochar would be 130 Pg CO2-Ce: These
results are possible at current levels of feedstock availability, while preserving
biodiversity, ecosystem stability and food security. They also show that conversion of all
sustainably obtained biomass to maximize bioenergy, rather than biochar, production can
offset a maximum of 10% of the current anthropogenic CO2-Ce emissions. The relative
climate-mitigation potentials of biochar and bioenergy depend on the fertility of the soil
amended and the C intensity of the fuel being offset, as well as the type of biomass.
Locations in which the soil fertility is high and coal is the fuel being offset are best suited
for bioenergy production. The climate-mitigation potential of biochar (with combined
energy production) is higher for all other situations.
IBI developed scenarios on carbon removal from the atmosphere by biochar. Those
scenarios primarily differ in the amount of biomass that was available in a sustainable way
from global Net Primary Production (NPP). The “Conservative” scenario assumes that
only biomass from cropping and forestry residues that otherwise has no use (about 27% of
the total residues) is available. The “Moderate” and “Optimistic” scenarios consider that
50% and 80%, respectively, of all cropping and forestry residues is available to produce
biochar. For each base scenario, IBI estimated the amount of biochar produced, as well as
the amounts of fossil fuel carbon emissions replaced by the energy generated during
biochar production. Moreover, IBI estimated the additional amount of carbon that could be
sequestered if CO2 emissions generated during biochar production were captured and
sequestered in the same way as proposed for coal combustion facilities.
18
The results of IBI scenarios show that the carbon sequestered in biochar can be 0.25 Gt
per year by 2030 in the “Conservative” scenario, and 1 Gt annually before 2050 in the
“Optimistic” scenario. An often-quoted analysis (Pacala and Socolow, 2004) shows a need
to have 7 Gt of carbon per year of reduced carbon emissions by 2054 just to keep
emissions at the 2004 level.
1.6.2. Biochar and soil
Biochar used as a soil amendment to improve soil fertility and plant growth has been
the focus of much research in the recent past (Zhang et al., 2012; Ibrahim et al., 2013). It
has shown promise as a sustainable amendment to enhance soil chemical properties
(Glaser et al., 2002; Lehmann et al., 2011). Soil may become degraded due to human
activities such as mining and industrial activities as well as the use of certain pesticides in
agriculture.
Because of its high organic C content, biochar has the potential to serve as a soil
conditioner to improve the physicochemical and biological properties of soils. Soil water
retention capacity increases with increase in organic C. About 18% increase in the water
holding capacity of soil containing biochar was reported (Glaser et al., 2002). Soil water
holding capacity is related to hydrophobicity and surface area of biochar, and the improved
soil structure following biochar application (Verheijen et al., 2010). Biochar amendments
have been reported to improve soil bulk density, porosity and hydraulic conductivity (Asai
et al., 2009; Jeffery et al., 2011; Abel et al., 2013). Moreover, a decrease of nutrient
leaching due to biochar application has been also reported (Sohi et al., 2009; Major et al.,
2010; Singh et al., 2010).
Biochar generally has a neutral to alkaline pH; however, acidic biochar has been also
reported (Chan et al., 2007). The pH of biochar depends on various factors including
feedstock type and the thermochemical process of production. The alkaline pH of biochar
induces a liming effect on acidic soils, thereby possibly increasing plant productivity. The
extent of liming effect of biochar depends on its acid neutralizing capacity that varies
depending on the feedstock and pyrolysis temperature. For example, biochar derived from
paper mill waste pyrolyzed at 550 °C has a liming value around 30% that one of CaCO3
(Zweiten et al., 2010). Significant increases in seed germination, plant growth, and crop
yields have been reported in the soils amended with biochars (Glaser et al., 2002).
19
The effect of biochar on microbial activity needs to be further investigated, especially
when considering the possibility of large applications of biochar in agronomic systems for
the purpose of increasing soil organic carbon. However, Lehmann et al. (2011) and more
recently Ameloot et al. (2013) reported direct and indirect interactive effects between
biochar and soil organisms. Although biochar does not provide a suitable habitat for soil
microorganisms (Quilliam et al., 2013), soil microbial activity may be indirectly
influenced by changes in the physicochemical properties, e.g. soil porosity, pH, cation
exchange capacity (CEC) and adsorption properties. In a direct way, microorganisms can
utilize a number of labile biochar constituents as an energy source (Cross and Sohi, 2011).
These are presumably either relatively untransformed biomass components that have not
been subjected to volatilization during pyrolysis (Ronsse et al., 2013) or volatilized
compounds that have recondensed in the biochar matrix during pyrolysis (Kloss et al.,
2012). However, Ameloot et al. (2014) reported that, in contrast to many short-term
laboratory studies, in field experiment biochar amendment led to a lowered or equal soil
microbial activity after 1-4 years incorporation in the field.
Some researches reported the potential role of biochar in reducing N losses. However,
to date little is known about the effects of biochar on the soil nitrogen (N) cycle. Yanai et
al. (2007) and Singh et al. (2010) have shown that biochar decreased N2O emissions
because of its ability to absorb water. In particular, Singh et al. (2010) hypothesized that
reduction of N2O emissions and ammonium leaching was determined by the increasing
biochar nutrient sorption capacity due to the higher oxidative reactions on its surface over
time.
Applying biochar together with organic or inorganic fertilizers can even enhance crop
yields (Lehmann et al., 2002). Studies show that when biochar is applied in soil, it
increases crop yield, reduces irrigation needs and enhances fertilizer efficiency (Steiner et
al., 2007; Blackwell et al. 2009). However, the biochar as a soil amendment for crop
production is still being investigated, and results so far are not conclusive. The application
of biochar to soils can boost crop yields by up to 60% or diminish yields by up to 30%,
mainly depending on the type of soil to which it is applied (Crane-Droesch et al., 2013).
Spokas et al. (2012), in their biochar review article, reported that biochar application rates
in research studies have range from <1 to over 100 t ha-1 and reported relative response to
biochar compared to the treatment that receives no biochar (0 t ha-1) from a reduction of 50
% to positive yield increases of ~200 %. Therefore, biochar applications affect crop yields
20
in highly variable ways. Such great variation likely stems from the large range of biochar
application rates, crops, and soil types used. In particular, the differences in results reflect:
1. Type of feedstock for biochar and the temperature and time of pyrolysis. The application
of different types of biochar can lead to very different responses (Rajkovich et al., 2012):
some types of biochar can increase crop production by over 100%, and others can reduce it
by a similar amount.
2. Differences in soil types. Positive effects on plant growth tend to be recorded from
highly- degraded and nutrient-depleted soils (Zwieten et al., 2009). Application of biochar
to fertile soils has not been shown to increase plant growth.
Therefore, the impact of biochar on the crop yield needs to be investigated further. In
fact, the extent with which biochar application might increase agricultural production is an
important driver in any attempt to develop systems that economically incorporate pyrolysis
products within the soil. It is not the only consideration (carbon sequestration is also very
important), but it requires long-term investment in agricultural experimentation.
1.6.3. Biochar and waste management
Biochar has great potential for managing the waste stream originating from animals or
plants; thus, decreasing the associated pollution loading to the environment. The use of
waste biomass for biochar production is not only economical but also beneficial. Making
biochar from biomass waste materials should create no competition for land with any other
land use option
such as food production or leaving the land in its pristine state.
Therefore, the conversion of wastes into biochar through pyrolysis is potentially an
effective waste management solution and economic feasibility.
Waste biomass that has been used to produce biochar includes crop residues (both field
residues and processing residues such as nut shells, fruit pits, bagasse, etc), forestry waste,
animal manure, food processing waste, paper mill waste, municipal solid waste and sewage
sludge (Cantrell et al., 2012; Enders et al., 2012).
Large amounts of agricultural, municipal, and forestry biomass are currently burned or
left to decompose and release CO2 and methane back into the atmosphere. They also can
pollute local ground and surface waters
a large issue for livestock wastes. Using these
materials to make biochar not only removes them from a pollution cycle, but biochar can
21
be obtained as a by-product of producing energy from this biomass. Moreover, pyrolyzing
the waste biomass, particularly animal manure and sewage sludge, kills any microbes
present, thereby reducing the environmental health effects (Lehmann and Joseph, 2009).
However, the persistence of toxic heavy metals in biochar developed from sewage sludge
and municipal solid waste (Lu et al., 2012) must be carefully handled before long-term
application to soils.
The annual worldwide production of wheat straw as agricultural waste was estimated to
be approximately 540 million tons in 2007 (Reddy and Yang, 2007). The straw might be
left on the field, burned, fed animals or used as industrial raw materials. As lignosulfonate
is the main component of paper mill waste, huge amount of lignosulfonate was generated
and the disposal of waste (liquid, solid and suspended matter) generated during the paper
manufacturing process contributed to a very high impact on the environment, but less of
them were utilized. Moreover large proportion of waste has been disposed of by burning
and discharging, resulting in not only a waste of resource but also a serious environmental
problem. Conversion of straw and lignosulfonate into biochar through pyrolysis has further
advantages of energy and environment.
1.6.4. Biochar and other environmental effects
Biochar not only improves chemical and biological soil properties but also can help
mitigate environmental issues by reducing of the mobility of heavy metals (Cu and Zn)
(Hua et al., 2009) and other organic soil contaminants (i.e. insecticides, Hilber et al.,
2009). The adsorption behavior of biochar for different contaminants (i.e., heavy metals,
organic pollutants and other pollutants) are different and well correlated with the properties
of contaminants. In addition, the adsorption mechanism may also depend on biochar’s
various properties including surface functional groups, specific surface area, porous
structure and mineral components.
One of the characteristics of biochars is possessing large surface areas, which implies a
high capacity for complexing heavy metals on their surface. Surface sorption of heavy
metals on biochar has been demonstrated on multiple occasions using scanning electron
microscopy (Beesley and Marmiroli, 2011; Lu et al., 2012). This sorption can be due to
complexation of the heavy metals with different functional groups present in the biochar,
due to the exchange of heavy metals with cations associated with biochar, such as Ca+2 and
22
Mg+2 (Lu et al., 2012), K+, Na+ and S (Uchimiya et al., 2011), or due to physical
adsorption (Lu et al., 2012). Also oxygen functional groups are known to stabilise heavy
metals in the biochar surface, particularly (Uchimiya et al., 2011) for softer acids like Pb+2
and Cu+2. In addition, Méndez et al. (2009) observed that Cu+2 sorption was related to the
elevated oxygenated surface groups, elevated superficial charge density and Ca+2 and Mg+2
exchange content of biochar. Furthermore, the surface area and porous structure of biochar
can also have effects on the adsorption of heavy metals. However, as the literature data
reported, the surface area and porous structure of biochar seem to have less effect on heavy
metal adsorption than oxygen-containing functional groups (Ding et al., 2014). Sorption
mechanisms are also highly dependent on soil type and the cations present in both biochar
and soil. Some other compounds present in the ash, such as carbonates, phosphates or
sulphates (Cao et al., 2009; Park et al., 2013) can also help to stabilise heavy metals by
precipitation of these compounds with the pollutants. Alkalinity of biochar can also be
partially responsible for the lower concentrations of available heavy metals found in
biochar-amended soils. Higher pH values after biochar addition can result in heavy metal
precipitation in soils. Biochar pH value increases with pyrolysis temperature (Wu et al.,
2012), which has been associated with a higher proportion of ash content (Cantrell et al.,
2012). Biochar can also reduce the mobility of heavy metals, altering their redox state of
those (Choppala et al., 2012). As an example, biochar addition could lead to the
transformation of Cr (VI) to the less mobile Cr (III) (Choppala et al., 2012). Therefore, the
possible adsorption mechanisms usually involved integrative effects of several kinds of
interactions including electrostatic attraction, ion exchange, physical adsorption, surface
complexation and/or precipitation (Fig. 4a). However, the relative contribution of the
different mechanisms to heavy metal immobilisation by different biochar remains
unknown.
Fellet et al. (2011) tried to use biochar to remediate a multicontaminated mine soil.
Biochar addition did not result in the decrease of the total heavy metal content of the soil;
however, biochar addition reduced the bioavailability of Cd, Pb and Zn and the mobility
(measured using a leaching experiment) of Cd, Cr and Pb. Uchimiya et al. (2012) analysed
the effects on soil heavy meals concentrations of 10 biochars prepared from 5 feedstocks at
2 different temperatures. They observed that manures with a high or low proportion of ash
or P were less effective to immobilise heavy metals. In contrast, biochars prepared at 700
°C were more effective, which could be attributed to transformations in the material,
including the removal of nitrogen containing heteroaromatic and leachable aliphatic
23
functional groups. They found Cu and Pb relatively easy to stabilise in soil, while Cd and
Ni response depended strongly on the type of biochar added to the soil.
Biochar application can also reduce the availability of organic contaminants such as
phenols in the soil (Gundale and DeLuca, 2007). The sorptive capacity of biochar to
organic contaminants in soil is controlled by carbonised and non-carbonised fractions and
the surface and bulk properties of biochar (Obst et al., 2011). Sorptive characteristics can
equally be affected by hydrophilic groups on biochar (James et al., 2005). The adsorption
mechanisms by which organic contaminants bind to biochars were also combined with
different kinds of interactions. In general, electrostatic interaction, hydrophobic effect,
hydrogen bonds, and pore-filling may be the main mechanisms for the adsorption of
organic contaminants onto biochar. The various mechanisms proposed for the interaction
of biochar with organic contaminants are summarized in Fig. 4b. For instance, the
adsorption of aromatic molecules such as PAHs to wood biochars is rapid and is assisted
by π-π electron interactions and pore-filling mechanisms (Chen et al., 2009), multilayer
adsorption, surface coverage, condensation in capillary pores, and adsorption into the
polymetric matrix (Werner et al., 2005). The results of different studies collected suggests
that electrostatic attraction was the dominant mechanism for adsorption of organic
contaminants onto the chars, with others performed as a contributing adsorption
mechanism (Inyang et al., 2014).
24
Figure 4. Summary of proposed mechanisms for (a) heavy metals and (b) organic
contaminants adsorption on biochars (Tan et al., 2015).
Recently, the biochar has been investigated for its effectiveness in saline soil
remediation. An interesting short incubation experiment (Wu et al., 2014) showed which
biochar can play a more important role in saline soil remediation reducing exchangeable
sodium percentage of saline soil and that biochar can improve soil fertility due to the
25
increment of soil organic carbon, cation exchange capacity and enhanced available
phosphorus.
1.7. Stability of biochar
The stability of biochar is of fundamental importance in the context of biochar use for
environmental management for two primary reasons: first, stability determines how long
carbon applied to soil, as biochar, will remain in soil and contribute to the mitigation of
climate change; second, stability will determine how long biochar will continue to provide
benefits to soil, plant, and water quality (Lehmann et al., 2006). It is well known that a
variable component of the carbon in many biochars is degradable on annual to decadal
timescales and hence, only a proportion of total carbon in biochar provides long-term
carbon sequestration (Bird et al., 1999; Zimmermann et al., 2012).
An increasing number of studies suggests that biochar can be degraded, by both biotic
and abiotic processes (Hamer et al., 2004; Cheng et al., 2008; Guggenberger et al., 2008).
However, in most of the studies the stability of biochar was assessed during laboratory
incubations, with fresh biochars added to soil (Zavalloni et al., 2011; Ameloot et al., 2013).
The duration of these experiments ranges from several weeks (Cross and Sohi, 2011) to
several years (Kuzyakov et al., 2009; Kuzyakov et al., 2014), allowing to understand
biochar stability under controlled laboratory conditions. On the contrary, there are only
few studies estimating biochar degradation rates in soil (Kuzyakov et al., 2009) and the
long-term stability of biochar in soils. This is because the changes of biochar content are
too small for any practical experimental period. Many studies estimating the
decomposition rates of biochar in soil are based on changes of CO2 efflux after biochar
application. This approach is unsuitable to estimate biochar decomposition because of the
much higher contribution of soil organic matter and plant residues mineralization of the
CO2 compared to biochar.
The complexity and chemical heterogeneity of biochar has made it difficult to establish
a single method suited to assessing the potential stability (Hammes et al., 2006) and hence,
there is no globally established method for determination of absolute stability for biochar.
However, a number of methods for comparing the relative stability of different biochar
materials have emerged. These include proximate analysis using the fixed carbon as a
measure of stability (ASTM Standard D3175; 2007), thermal analysis (thermogravimetry,
26
TG; de la Rosa et al., 2008), molecular markers by means of pyrolysis-gas
chromatography-mass spectrometry (Py-GC-MS; Kaal et al., 2008, 2009, Conti et al.,
2014), benzene polycarboxylic acid method (Brodowski et al., 2005), O:C or H:C molar
ratios (Spokas, 2010; Enders et al., 2012; IBI Guidelines, 2012) and chemical oxidation
(Cross and Sohi, 2013). Further information on the studies conducted on biochar stability
can be found in the introductory section of chapters 5.1 and 5.2.
1.8. Biochar and pollutants
Biochar quality guidelines have been recently proposed such as the IBI Biochar
Standard (IBI, 2013), the European Biochar Certificate (EBC, 2014) or the UK Biochar
Quality Mandate (BQM, Shackley et al., 2013). In these standards, environmental risks are
accounted for by the inclusion of limit values for physicochemical properties, including
pollutants such as heavy metals, dioxins/furans,
and polychlorinated biphenyls (PCBs). For instance, the IBI sets a range of maximum
allowed threshold values (varying between different countries) for the sum of the 16 US
Environmental Protection Agency’s (EPA) PAHs in biochar to 6-20 mg kg-1 dry weight
(IBI, 2013). Similarly, the EBC requires PAHs to be below 4 and 12 mg kg-1
dw,
(dw)
in
premium and basic grade biochars, respectively (EBC, 2014).
Research shows (Hale et al., 2012 and Oleszczuk et al., 2013) that biochar can contain
dangerous inorganic contaminants (heavy metals) and organic ones (e.g. polycyclic
aromatic hydrocarbons (PAH) as well as dioxins and furans (PCDD/Fs)). The presence of
contaminants, therefore, poses a question mark over the common utilisation of biochars,
especially for the amendment of soils used for crop plant cultures. In the case of high
levels of contaminants there is the risk of their uptake by plants or migration down the soil
profile to groundwaters. This may have negative effects for humans, for the environment
and for living organisms. Among the threats, most frequently mentioned is the
contamination of biochar with PAHs and heavy metals. While in the case of heavy metals
their levels are at relatively low values (Freddo et al., 2012) and depend on the content of
trace metals in the initial material, studies concerning PAHs indicate (Freddo et al., 2012;
Hale et al., 2012; Hilber et al., 2012; Keiluweit et al., 2012; Oleszczuk et al., 2013; Fabbri
et al., 2013) that biochars may be contaminated with those compounds to a significant
degree. PAHs are formed during the pyrolysis of organic matter (including biomass) and
27
the content in biochars varies with relation to the feedstock, and also to the conditions of
the pyrolysis process (Manyà, 2012; Wiedner et al., 2013).
The application of biochar (containing high levels of PAHs) to soils, even at small
doses, may undeniably cause an increase in the soil content of those contaminants.
Moreover, s
PCDD/Fs could be present in the biochars produced from feedstock that contain
chlorine. Biochar feedstocks such as grasses, straws and food waste (which contains
sodium chloride, i.e., salt) can be a source of chlorine. Other potential sources of chlorine
in biochar feedstocks include biomass that has been exposed to salt (such as crops or trees
grown near seashores), and the biomass fraction of municipal solid waste that may be
contaminated with polyvinyl chloride (PVC) or other chlorine-containing plastics.
Research concerning dioxin indicates that biochars may be contaminated with those
pollutants (Downie et al., 2011). Moreover, in biochars from food waste have been found
relatively high concentration of dioxin likely due to the high salt (sodium chloride) content
in food waste. However, in the biochars studied the total dioxin concentrations (0.005-1.20
ng Kg-1) (Hale et al., 2012) are lower than the guideline values for dioxin and furans in
biochar of the EBC (20 ng Kg-1 TEQ) and IBI (17 ng Kg-1 TEQ). Further information on
the PAH topic has been reported in the introductory part of section 3.
1.9. Biochar: some non-negligible issues
Owing to the extensive range of combinations of biochar, soils and plants, much
research still needs to be undertaken to understand the large variety of resulting
interactions and their effects. As research progresses, it will be possible to make
28
extrapolations with increasing robustness as, for example, the database upon which metaanalysis can be performed grows. Such information is vital to guide the development of
certification schemes such as that proposed by the IBI, and the EBC, which is already
implemented in part of Europe, as well as to guide policy.
Interaction of biochar with soil microbial communities and plants
The physical, biological and chemical processes that biochar may exert on microbial
communities and their symbiotic interaction with plants, and possibly enhanced nutrient
use efficiency, are not yet understood. The apparent contradiction between the high
stability of biochar, soil organic matter accumulation and apparent enhancement of soil
microbial activity needs to be resolved. In the future work, the effects of biochar on
various soil biota groups, their diversity and functioning need to be carefully considered.
Moreover, further research biochar needs to involve a careful selection of the feedstock
and pyrolysis conditions to find an optimal match of biochar type to the intended
ecosystem goal.
Biochar erosion, transport and fate
The loss of biochar through vertical or lateral flow is not quantified, and only recently
have studies been initiated to examine movement through soil profiles and into water
ways. It should be noted however that transport of biochar through the profile does not
impact on its direct carbon sequestration potential.
Biochar stability
A key requirement for the use of biochar as tool for environmental management is that
the carbon in the biochar is stable, meaning that a substantial fraction of the carbon
sequestered is not re-mineralized on at least centennial timescales. However, a variable
component of the carbon in many biochars is degradable on annual to decadal timescales
and hence, only a proportion of total carbon in biochar provides long-term carbon
sequestration. Although our understanding of biochar carbon stability has improved in
recent years, there is limited research on process conditions to produce a biochar suitable
and highly stable for the long-term carbon sequestration.
Pollutants environmental fates
The fate of contaminates in the environment is of prime importance in order to prevent
severe contamination to the environment. The environmental fates of biochar-associated
pollutions added to soil are still poorly understood. In particular, further research work is
29
required to improve understanding of the role biochar plays in sorbing PAHs and on
microbial activity and how this influences the concentration of PAHs in soil and their
persistence in the environment.
Synergistic effects
The interactions of biochar with soil organic matter as well as the mineral matrix need
to be assessed in order to determine the nature and the environmental conditions under
which synergistic effects develop.
Water holding capacity
The contribution that biochar can make to water retention, macro-aggregation and soil
stability is poorly understood – yet should be of critical importance in climate change
adaptation, where mitigating drought, nutrient loss and erosion are critical.
Cation exchange capacity (CEC)
While the CEC of fresh char itself is not very high biochar that has resided in soil for
hundreds of years has been shown to have much higher CECs, comparable to those of
zeolites. However, several studies have reported an increase in soil CEC after the
application of fresh biochar. Thus, the processes that are instrumental in developing CEC
over time as well as the effects that lead to an increase in CEC by addition of fresh (low
CEC) biochar requires detailed understanding.
Decreased emissions of non-CO2 greenhouse gases (e.g. N2O and CH4)
The currently available data on the effect of biochar additions on trace gas emission is
very limited, but has a potentially great impact on the net benefit of biochar application.
Therefore, further research work is required to determine the impact of biochar on the
emission of N2O and CH4.
Soil carbon modelling
Modelling of the linked carbon and nitrogen cycles in soil with and without application
of biochar is essential to understanding the fundamental mechanisms referred to above, and
the impact on soil-based emissions of greenhouse gases.
Project specific Life Cycle Assessment (LCA)
The total environmental life cycle assessment has been conducted for some biochar
case studies. Greenhouse balances, for example, are very project specific and hence there
30
is opportunity to assess the benefits over a large range of feedstock, process and biochar
application scenarios.
31
2. Aim of the thesis
Studies on biochar are relatively recent, leaving several aspects unexplored or not fully
developed. Further research on the impact of biochar in the environment in both the long
and the short term is required both to avoid unforeseen consequences and to provide
evidences of further potential benefits. In particular, the environmental potential and
limitation of biochar in soil applications requires a full understanding of the stability and
fate of carbon fractions and trace contaminants, in particular PAHs. For this reasons this
thesis was focused on the assessment of biochar stability and the occurrence and fate of
PAHs.
Regarding PAHs, biochar is the by-product of a thermochemical process. Therefore,
the formation of PAHs from biomass pyrolysis and their occurrence in biochar is inevitable
and must be considered and properly evaluated in order to avoid or limit occupational
exposure, land contamination and PAHs transfer to crops. Due to the carbonaceous nature
of biochar that has a great affinity for polyaromatic compounds, the analysis of PAHs is
challenging, no certified reference materials are available, and standardised methods are
being developed. In order to determine the level of PAHs with reliable analytical
procedures and evaluate the potential negative impact, this thesis aimed at:

developing a well characterized analytical method for the determination of
PAHs in pure biochars as well as in soil-biochar matrices.

measuring the levels of PAHs in biochars from different feedstock and process
conditions, searching for causal relationships, extending the analysis to EUPAHs (food safety) along with EPA-PAHs (environmental protection).

assessing the long-term impacts of biochar additions, at different applications
rates, on PAHs concentration in agricultural soils;

evaluating their possible role in the phytoxicity of animal vs. plant derived
biochar.
Regarding the environmental stability, this is probably the most crucial and less known
among the properties of biochar of interest for assessing its benefits to soil organic carbon
and CO2 mitigation. In this topic, the thesis was focused to:
32
I.
The determination and quantification of labile and resistant carbon fractions in
biochar by hydropyrolysis (HyPy).
II.
The molecular characterisation of the biochar labile fraction by HyPy combined
with GC-MS.
III.
The assessment of the impact of production conditions on biochar stability by
pyrolysis-GC/MS and HyPy.
IV.
The identification of the resistant carbon fractions and the characterization of
the labile organic carbon in biochar amended soils in a four years field study.
33
3. Determination of PAHs: method development and application
3.1. Determination of polycyclic aromatic hydrocarbons in biochar and biochar
amended soil
3.1.1. Introduction
Biochar is a co-product from biomass pyrolysis that is targeted as a material with
applications in environmental and agricultural management, as well as a vehicle for carbon
sequestration (Sohi et al., 2010; Manyà et al., 2012). As the interest toward biochar is
steeply growing, safety procedures for ensuring human health and preservation of the
environment are imperative. Polycyclic aromatic hydrocarbons (PAHs) are well known
carcinogenic and persistent pollutants that are ubiquitous in the environment. PAHs are
formed during the pyrolysis of biomass (Fabbri et al., 2010) and their occurrence in
biochar (Hale et al., 2012; Hilber et al., 2012; Schimmelpfennig and Glaser., 2012;
Keiluweit et al., 2012; Freddo et al., 2012; Kloss et al., 2012) along with its possibly
released into the environment need to be addressed. PAH production has also been
confirmed during the production of charcoal by pyrolysis (Ré-Poppi et al., 2002; Mara Dos
Santos Barbosa et al., 2006) and natural wildfires (Kim et al., 2003). Human exposure of
PAHs might occur through different pathways, such as inhalation of particles generated
during synthesis, handling and field applications of biochar or the ingestion of
fruit/vegetables grown in biochar amended soil. Therefore, determining the content of
PAHs in biochar is of utmost importance to establish risk assessment of biochar usage.
The worldwide distribution of PAHs in soils span over five orders of magnitude and is
related to source (atmospheric input) and sorption ability of soil organic matter and black
carbon (Nam et al., 2009). The inclusion of carbonaceous residues in soil could increase
PAHs sorption on humic matter (Cornelissen et al., 2005; Oen et al., 2006; Poerschmann et
al., 2007; Brandli et al., 2008) and biochar (Hale et al., 2011; Oleszczuk et al., 2012). In
this respect, soil application of biochar might represent a source and/or a sink of PAHs. All
these aspects need to be considered when dealing with the origin of PAHs in soil amended
with biochar.
A reliable methodology of PAH analysis is a first requisite towards risk assessment.
Recent studies have examined the content of PAHs in biochar (Hale et al., 2012; Hilber et
al., 2012; Keiluweit et al., 2012). These results have provided a comprehensive picture on
34
the levels and availability of PAHs in biochar (Hale et al., 2012), the influence of pyrolysis
temperature (Keiluweit et al., 2012), as well as critical aspects of validation (Hilber et al.,
2012). Analytical methods described in these studies have utilized toluene as the extracting
solvent. In fact, it was demonstrated that toluene is superior to other solvents for
carbonaceous materials (Jonker et al., 2002). Nonetheless, extraction efficiencies are not
always quantitative, especially in the case of low molecular weight (LMW) PAHs. In
particular, naphthalene is problematic because of the high boiling point of toluene (111°C)
which causes the loss of semi-volatile PAHs during the preconcentration step (Hilber et al.,
2012; Keiluweit et al., 2012). Naphthalene is considered a possible carcinogenic to humans
(IARC group 2B) and genotoxic to plants (Aina et al., 2006). Incidentally, naphthalene is
often the most abundant PAH in biochar (Hale et al., 2012; Hilber et al., 2012;
Schimmelpfennig and Glaser, 2012; Freddo et al., 2012; Kloss et al., 2012; Spokas et al.,
2011). Naphthalene and its isotopically labelled version are often employed in studies
aimed at investigating the fate of PAHs in the environment (Wild et al., 1994; Fraser et al.,
1998; Kipopoulou et al., 1999; Motelay-Massei et al., 2006). In general, LMW PAHs are
absorbed at higher rates than high molecular weight (HMW) PAHs (Kipopoulou et al.,
1999; Motelay-Massei et al., 2006; Tao et al., 2004), and naphthalene presence could
affect the growth/response of the soil microbial community (Loibner et al., 2004; Krang et
al., 2007).
Although present at lower concentrations, HMW PAHs pose the highest health and
environmental hazards due to the established carcinogenic potential of this class of
compounds. Because of biochar’s proposed use in crops and potential human exposure of
biochar PAHs through bioaccumulation in agricultural products, biochar sorbed PAH
concentrations could be a matter of concern (Ahn et al., 2008; Meudec et al., 2006; ReySalgueiro et al., 2009). On the basis of their occurrence and carcinogenicity, 15PAHs have
been identified as priority hazardous substances in food by the European Union (EU)
(ECR, 2006)] and 16 PAHs by US Environmental Protection Agency (USEPA) (2002), 8
of them are shared across both lists. While studies have been reported on the occurrence of
USEPA PAHs in biochar due to the widespread inclusion of these compounds in
worldwide environmental legislation, very limited information is available on the
occurrence of EU PAHs on biochar.
In addition, recent studies were focused on the analysis of PAHs in solely biochar, but
the robustness of the solvent extraction method to extract PAHs when biochar is present in
the soil was not fully investigated. It is important that a method developed for the analysis
35
of solely biochar should be equally accurate for the biochar-soil matrix. In this context, the
use of (cyclo)hexane/acetone mixtures as an extracting solvent in PAH determination in
soil is rather common (e.g. Gfrerer et al., 2002; Shu et al., 2003; Beesley et al., 2010). In
fact, a relatively polar solvents like acetone has been cited as beneficial for the extraction
of hydrophobic PAHs from soil (Pena et al., 2007).
The present study is aimed at developing a well characterized method for the
determination of PAH in biochars and soils amended with biochar by GC–MS. To this
purpose, several solvent and extraction procedures were examined using the 16 EPA PAHs
as targeted PAHs on a biochar utilized in agronomic field studies (Fellet et al., 2011). The
method was then applied to a set of biochars investigated as soil amendments of different
origin and from different process conditions (Fabbri et al., 2012). Besides the EPA PAHs,
the level of EU PAHs in these biochars was investigated as well.
3.1.2. Materials and methods
3.1.2.1. Reagents and standards
Cyclohexane, acetone, acetonitrile, dichloromethane, toluene, ethyl acetate (all ultrapurity), and surrogate standard mix (for USEPA 525) containing acenaphthene-d10,
phenanthrene-d10 and chrysene-d12 at concentrations of 500 mg l-1 each in acetone were
purchased from Sigma-Aldrich. PAH-Mix solution containing naphthalene, acenaphtylene,
acenaphthene, fluorene, phenanthrene, anthracene, fluoranthene, pyrene, chrysene,
benzo[a]anthracene,
benzo[b]fluoranthene,
benzo[k]fluoranthene,
benzo[a]pyrene,
dibenz[a,h]anthracene, indeno[1,2,3-cd]pyrene and benzo[ghi]perylene certified at
concentrations of 10 mg l-1 for each species in acetonitrile was purchased from Sulpeco
(Belleforte, PA, USA).
PAH-Mix standards in acetonitrile (10 mg l-1) of EU PAHs were obtained from Dr.
Ehrenstorfer GmbH (Augsburg, Germany): benzo[a]anthracene, benzo[b]fluoranthene,
benzo[j]fluoranthene,
chrysene,
benzo[k]fluoranthene,
cyclopenta[c,d]pyrene,
benzo[ghi]perylene,
dibenzo[a,h]anthracene,
benzo[a]pyrene,
dibenzo[a,e]pyrene,
dibenzo[a,h]pyrene, dibenzo[a,i]pyrene, dibenzo[a,l]pyrene, indeno[1,2,3-c,d]pyrene, 5methylchrysene. Standard mix solutions containing the 15 PAHs at concentrations of 1 mg
l-1 were prepared in acetone/cyclohexane (1:1, v/v) and stored at room temperature in the
dark.
36
A solution of 1,3,5-tri-tert-butylbenzene (TTB, 12.7 mg l-1) in acetone:cyclohexane
(1:1, v/v) was prepared by weighing the pure compound purchased from Sigma-Aldrich.
3.1.2.2. Soil and biochar samples
A natural matrix soil certified reference material ERM–CC013a (manufactured by
Federal Institute for Materials Research and Testing; Berlin, Germany) containing 15
PAHs with concentrations ranging from 1.14 to 12.9 mg kg-1 was used for the validation of
the method for soil. An internal reference biochar sample (here named as reference
biochar, or RB) was utilized for method optimization. This was a commercially available
biochar created by the slow pyrolysis of orchard pruning, which was kindly provided by
the Department of Agriculture and Environmental Sciences (DISA) University of Udine
(Fellet et al., 2011). This reference biochar was homogenized and then mixed with an
agricultural soil (dried and sieved 2 mm) at a 1.16% (w/w) amendment level. This
concentration corresponded to an application of 36 t biochar ha-1 (assuming a soil with 1.2
g cm-3 density and 0.3 m depth) (Schimmelpfennig and Glaser., 2012; Zavalloni et al.,
2011), which is within the range currently investigated for biochar use in agriculture (20–
60 t biochar ha-1) (Baronti et al., 2010).
Additional biochars evaluated were from an on going study on the impact of biochar
additions on greenhouse gas production potentials conducted by the USDA-ARS Biochar
and Pyrolysis Initiative. The full characterization of these biochars (i.e. ultimate and
proximate analysis, Py-GC–MS, and microbial CO2 production) was reported in a previous
publication [40]. This group provides across-section of currently available biochars for
agricultural field applications.
3.1.2.3. Sample treatment
3.1.2.3.1. Optimized sample pretreatment: soxhlet extraction and clean up
About 1 g of biochar (or 5 g soil sample) was placed into the extraction cellulose
thimble, spiked with 0.1 ml of surrogate standard mix (Supelco for EPA 525 containing
acenaphthene-d10, phenanthrene-d10 and chrysene-d12 5 µg ml-1 each in acetonitrile). The
thimble was covered with cotton wool, and inserted into the Soxhlet extractor. Soxhlet
extraction thimbles (and the Soxhlet apparatus) were pre-cleaned by a 4 h Soxhlet
extraction with acetone/cyclohexane (1:1, v/v). Extraction was carried out with 160 ml of
37
extraction solvents (acetone/cyclohexane (1:1, v/v)) mixture for 36 h (4 cycles h-1). The
Soxhlet apparatus was covered with an aluminum foil to avoid exposure to daylight, which
prevents PAH photodegradation. The extraction solvent was filtered, added with 1 ml of nnonane, and then carefully evaporated by rotatory vacuum evaporation at 40 °C. The
concentrated extract was collected and loaded onto a silica gel cartridge (6 ml, 1 g DSC-Si
Supelco washed with ethyl acetate, dried and conditioned with 4 ml cyclohexane). After
purification with 1 ml of cyclohexane, PAHs were eluted with 4 ml of acetone/cyclohexane (1:1, v/v). The obtained solution was then blown down to 10–50 µl
under nitrogen and spiked with 10 µl of the internal standard solution (TTB at 12.7 mg l-1)
prior to GC–MS analysis.
3.1.2.3.2. Reflux extraction
Four different solvent systems (toluene, dichloromethane, acetone:cyclohexane 1:1
(v/v) and acetone:cyclohexane 1:5 (v/v)) were compared by means of reflux extraction. To
this purpose PAHs were extracted from the biochar (2 g reference biochar added with 0.1
ml of surrogate standard mix) by refluxing for 4 h with 80 ml solvent. The extract was
filtered and concentrated to ∼100 µl by using rotary evaporator and then under a nitrogen
stream. The obtained solution was spiked with 10 µl of internal standard (12.7 mg l -1 TTB)
and then analyzed by GC–MS.
3.1.2.3.3. Ultrasonication extraction
Each homogenized reference biochar sample (1 g) was transferred into a Pyrex tube, and
20 ml of acetone/cyclohexane (1:1, v/v) were added. The sample was ultrasonicated for 30
min with occasional swirling. The extraction solutions were then centrifuged and the
supernatant filtered into a 50 ml beaker using a 9.0 cm GF/C glass microfibre filter
(Whatman International, Maidstone, UK). The obtained solutions were reduced to 2 ml
using a rotary evaporator and transferred into 4 ml vials. These solutions were further
reduced using nitrogen gas, spiked with 10 µl of 12.7 mg l-1 TTB, and analyzed by GC–
MS.
38
3.1.2.4. GC–MS
GC–MS analyses were performed using a 6850 Agilent HP gas chromatograph
connected to a 5975 Agilent HP quadrupole mass spectrometer. Analytes were separated
by
a
HP-5MS
fused-silica
capillary
column
(stationary
phase
poly[5%
diphenyl/95%dimethyl]siloxane, 30 m × 0.25 mm i.d., 0.25 mm film thickness), using
helium as the carrier gas. Samples (1 µl) were injected under splitless conditions (1 min,
then split ratio 1:50 to the end of analysis) with an injector temperature of 280 °C. The
following thermal program of the capillary column was used: 50 °C to 100 °C at 20 °C
min-1, then from 100 °C to 300 °C at 5 °C min-1, then a hold for 2.5 min at 300 °C. The
mass spectrometer operated under electron ionization (70 eV) and acquisition was
performed on single ion monitoring (SIM) at the molecular ion of each PAH at the time
windows corresponding to the elution region of the target PAH. Acenaphthene-d10 was
utilized
to
quantify
naphthalene,
acenaphthylene,
acenaphthene
and
fluorene;
phenanthrene-d10 to quantify phenanthrene, anthracene, fluoranthene and pyrene;
chrysene-d12 to quantify the remaining PAHs. Quantitation of EPA PAHs was based on the
calibration curve (Section 3.1.2.5), while in the case of EU PAHs a single point calibration
(1 mg l-1, Section 3.1.2.1) was utilized.
3.1.2.5. Method validation
The figures of merit were reported for the EPA PAHs. Recovery of surrogated PAHs
was determined with respect to the internal standard TTB. The procedural blank
concentrations were determined as the average of five empty thimble runs. Procedural
blanks were run periodically. Precision of the procedure was determined by four replicate
analyses of reference biochar sample. Calibration was performed in the 0.0025–1.25 mg l-1
interval by serial dilutions of the 10 µg ml-1 EPA PAH calibration mix (Supelco). Three
replicates were performed at each concentration level and the resulting instrumental
response was homoscedastic for each PAH ( = 0.05, Cochran test), therefore the leastsquares regression line was utilized for quantification (R2 values were 0.993–0.999). Limit
of detection (LOD) and limit of quantification (LOQ) were estimated for each analyte by
using Eqs. (1) and (2)
39
LOD = 3 sb / a
(1)
LOQ = 10 sb / a
(2)
Table 3.1.1. Limits of detection (LOD), limits of quantification (LOD), mean
concentration of EPA PAHs in reference biochar (RB) and relative standard deviations
(RSD) from four replicates.
PAH
Naphthalene
Acenaphtylene
Acenaphthene
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
Chrysene
Benzo[a]anthracene
Benzo[b]fluoranthene
Benzo[k]fluoranthene
Benzo[a]pyrene
Indeno[1,2,3-cd]pyrene
Dibenzo[a,h]anthracene
Benzo[ghi]perylene
LOD (ng g-1)
LOQ (ng g-1)
RB (μg g-1)
RSD (%)
0.08
0.01
0.03
0.03
0.4
0.03
0.08
0.06
0.1
0.08
0.2
0.09
0.2
0.2
0.3
0.1
0.2
0.03
0.1
0.1
1
0.1
0.3
0.2
0.4
0.3
0.5
0.3
0.8
0.7
0.9
0.4
1.75
0.026
0.034
0.071
0.71
0.13
0.30
0.35
0.095
0.095
0.13
0.10
0.19
0.15
0.056
0.15
8
13
5
10
12
13
11
11
9
9
6
18
14
16
15
8
where sb stands for the mean standard deviation of peak areas integrated at the retention
time of the PAH from procedural blanks and a for the slope of the calibration curve.
Results of LOD, LOQ and precision (%RSD) are listed in Table 3.1.1.
40
3.1.3. Results and discussion
3.1.3.1. Solvent selection
The choice of the extracting solvent is a crucial parameter in the analysis of PAHs in
carbonized materials (soot, charcoal) because hydrophobic contaminants are tightly bound
to the aromatic matrix (Jonker et al., 2002). In this study, the extraction ability of four
different solvent systems was preliminary evaluated by means of reflux extraction under
the same conditions. Toluene, solely (Hale et al., 2012; Hilber et al., 2012) or mixed with
methanol (Keiluweit et al., 2012), was the solvent of choice in the determination of PAHs
in biochar reported in recent literature and therefore included in this comparison.
Dichloromethane is a rather common solvent in the extraction of PAHs in several matrices,
including wood chars (Brown et al., 2007) and biochar (Freddo et al., 2012).
Acetone/hexane mixtures were described in the analysis of PAHs in charcoal and soot
samples (Jonker et al., 2002).
The recovery of surrogate PAHs for each extraction system is reported in Table 3.1.2.
Toluene is the best extracting solvent in the case of spiked d-phenanthrene and d-chrysene.
This finding is in agreement with previous studies showing the strong extraction efficiency
of toluene in comparison to other solvents and solvent/mixtures (Hilber et al., 2012; Jonker
et al., 2002). However, in the case of spiked d-acenaphthene, dichloromethane and
acetone/cyclohexane 1:1 exhibited higher extraction efficiency than toluene (83 and 80%
vs. 68%). The loss of LMW PAHs in the case of toluene was caused by the analytical
procedure following the extraction step, as blank analysis with toluene (resulting from
solvent evaporation) confirmed a recovery of 65 ± 11% of d-acenaphthene. A similar result
was reported by Hilber et al. (2012), who suspected a cross-contamination by naphthalene
possibly due to extended toluene removal. When examining the PAH concentrations as a
function of solvent, the detected concentrations of the LMW PAHs were the lowest with
toluene (0.84 µg g-1) and highest with acetone/cyclohexane 1/1 (1.37 µg g-1). Therefore,
the solvent mixture of acetone/cyclohexane was selected for the method optimization,
because of its superior extraction efficiency for naphthalene (the most common PAH
detected on biochar; see below), its widespread use in soil analysis of PAHs, and its
reduced toxicity compared to toluene and dichloromethane.
41
Table 3.1.2. Recovery of surrogate PAHs using different extraction procedures of
reference biochar (RB).
Acenaphthene-d10
recovery (%)
Phenanthrene-d10
recovery (%)
Chrysene-d12
recovery (%)
80
56
83
68
41
38
50
68
7
7
11
58
9
4
0.4
Soxhlet extraction (18 hours)
Acetone/cyclohexane 1/1
Acetone/cyclohexane 5/1
Acetone
75
76
84
66
37
58
29
10
29
Soxhlet extraction (36 hours)
Acetone/cyclohexane 1/1
88
77
67
Reflux extraction
Acetone/cyclohexane 1/1
Acetone/cyclohexane 1/5
Dichloromethane
Toluene
Ultrasonication extraction
Acetone/cyclohexane 1/1
3.1.3.2. Selection of the extraction procedure
The recovery of surrogate PAHs from reflux extraction with acetone:cyclohexane 1:1
were compared with Soxhlet extraction (18 h) and ultrasonic extraction (Table 3.1.2).
Ultrasonic extraction had very low recoveries (<10%) and therefore was not investigated
further. As expected, the recovery of d-chrysene by Soxhlet extraction increased with
respect to reflux conditions. Increasing (100%, v/v) or decreasing (20%, v/v) the mixing
ratio of acetone with respect to the 1:1 acetone:cyclohexane mixture (i.e. 50%, v/v) did not
significantly
improve
the
recovery
of
the
surrogate
PAHs.
Therefore,
the
acetone:cyclohexane mixture 1:1 was selected to investigate the effect of the extraction
time on the recovery. The results, depicted in Fig. 3.1.1, show that the higher recoveries
were achieved with longer extraction times, which is in agreement with a previous study
(Hilber et al., 2012). Interestingly, the same study showed that accelerated solvent
42
extraction (ASE) was a less efficient than Soxhlet extraction (Hilber et al., 2012).
However, prolonged extractions were problematic and did not guaranteed high recovery.
Figure 3.1.1. Recovery of deuterated PAHs vs. soxhlet extraction times with
acetone:cyclohexane 1:1 v/v of reference biochar (mean values and 1 s.d. from four
replicates).
We decided to focus on the behaviour of two HMW PAHs representative of five rings
(benzo[a]pyrene) and six (indeno[1,2,3,cd]pyrene) rings as the target compounds for
optimizing the extraction time. Their concentrations increased significantly when the
extraction time was increased from 18 to 36 h, after which time the concentration remained
almost constant. Thus, 36 h of extraction were selected for the final procedure.
43
3.1.3.3. Final procedure applied to reference biochar and soil
The final procedure was described in detail in Section 3.1.2.3.1. The USEPA PAH
concentrations of reference biochar are reported in Table 1 along with the relative standard
deviations. A typical chromatogram is presented in Fig. 3.1.2. The precision (expressed as
RSD from four replicates) was good, being within the 5–18% interval. The recoveries of
surrogate PAHs were satisfactory (67, 77, and 88% for d-acenaphthene, d-phenenthrene,
and d-chrysene, respectively, Table 3.1.2). This is also considered a good result
considering that PAHs are strongly associated to the aromatic carbonaceous matrix of
biochar. These results are on the higher end of PAH recoveries currently reported for
biochar materials. Hilber et al. (2012) reported 42–72% recovery range for several
deuterated PAHs (from d-naphthalene to d12-indeno[1,2,3-cd]pyrene), and similar values
(56–79%) were reported by Hale et al. (2012). The accuracy of the method developed for
biochar was tested on the soil matrix by the analysis of the certified soil (ERM-CC013a).
The difference between the mean measured and certified values (Table 3.1.3) were lower
than the expanded uncertainty of that difference for the majority of PAHs, attesting the
validity of the method for the soil matrix (Linsinger et al., 2005). Then, the ability of the
method to analyze PAHs in the biochar amended soil was evaluated. The obtained
concentrations of PAHs in the untreated soil and in the soil amended with biochar are
presented in Table 3.1.4. The total PAH concentration in the amended soil is significantly
higher than that in the untreated soil. In particular, the concentration of naphthalene is
0.0263 µg g-1 against 0.0098 µg g-1 in the untreated soil, a quite large difference due to
naphthalene being the most abundant PAH in biochar at 1.75 µg g-1. The excess
naphthalene in the treated soil of 0.0263 – 0.0098 = 0.0165 µg g-1 is slightly lower than
that expected from the quantity of naphthalene added with biochar corresponding to 1.75 ×
1.16% = 0.0203 µg g-1. Overall, the correspondence between the measured excess and
expected is (0.0165–0.0203)/0.0203 = -0.19 (or -19%), which is an acceptable result and
good demonstration of the accuracy of the method for LMW PAH compounds, which has
been a shortcoming of some of the existing methods [i.e. 5].
44
Figure 3.1.2. GC-MS (SIM) chromatogram obtained from the analysis of reference
biochar (RB). Peak numbers refers to PAHs listed in table 3.1.5.
CRYd
Counts
Counts
1
10
65000
55000
240000
45000
35000
200000
25000
160000
3
120000
80000
11
15000
9
ACEd
PHEd
32.00
5
36.00
TTB
7
40000
2
10.00
4
15.00
18
17 19
13 14
16
12
5000
40.00
44.00
Min 48.00
CRYd
8
6
20.00
25.00
30.00
35.00
40.00
45.00
Min
Table 3.1.3. Validation of the optimized method for the soil matrix through the analysis of
the certified material ERM – CC013a.
PAH
Naphthalene
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
Benzo[a]anthracene
Chrysene
Benzo[b]fluoranthene
Benzo[k]fluoranthene
Benzo[a]pyrene
Benzo[ghi]perylene
Indeno[1,2,3-cd]pyrene
Measured
concentration
(μg g-1)
Certified
value
(μg g-1)
Relative
error
(%)
2.2 ± 0.2
1.30 ± 0.11
12.4 ± 0.3
1.96 ± 0.09
12.0 ± 0.5
8.4 ± 0.6
5.1 ± 0.3
6.3 ± 0.3
6.4 ± 0.4
4.0 ± 0.4
4.6 ± 0.4
4.3 ± 0.7
5.5 ± 0.9
2.4 ± 0.5
1.14 ± 0.11
12.0 ± 0.6
1.41 ± 0.22
12.9 ± 0.7
9.6 ± 0.3
5.6 ± 0.5
5.3 ± 0.8
7.1 ± 1.0
3.4 ± 0.4
4.9 ± 0.7
4.6 ± 0.5
5.2 ± 1.0
-9
+13
+2
+32
-9
-15
-11
+15
-12
+14
-8
-8
+3
45
A similar calculation was performed for the other PAHs, and the results are reported in
last column of Table 3.1.4. These differences between the calculated and measured values
were satisfactory for the most abundant PAHs in biochar (Table 3.1.4; at the ±20% level).
These data support that the proposed method was capable to extract PAHs from a biochar
amended soil, a PAH contaminated soil, and the original biochar.
Table 3.1.4. Observed concentration of PAHs in an agricultural soil and a corresponding
biochar amended soil (1.16% (w/w) of reference biochar RB).
PAHs
Naphthalene
Acenaphtylene
Acenaphthene
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
Chrysene
Benzo[a]anthracene
Benzo[b]fluoranthene
Benzo[k]fluoranthene
Benzo[a]pyrene
Indeno[1,2,3-cd]pyrene
Dibenzo[a,h]anthracene
Benzo[ghi]perylene
Total
Soil
(µg g-1)
Soil + biochar
(µg g-1)
Difference from
expected
(%)
0.0098 ± 0.0002
n.d.
n.d.
0.0023 ± 0.0008
0.0118 ± 0.0036
0.0003 ± 0.0002
0.0035 ± 0.0010
0.0031 ± 0.0007
0.0007 ± 0.0003
0.0039 ± 0.0007
0.0067 ± 0.0014
0.0005 ± 0.0001
0.0001 ± 0.0002
0.0023 ± 0.0008
0.0009 ± 0.0002
0.0046 ± 0.0011
0.0506 ± 0.017
0.0263 ± 0.0046
n.d.
n.d.
0.0033 ± 0.0006
0.0212 ± 0.0063
0.0014 ± 0.0014
0.0075 ± 0.0030
0.0069 ± 0.0020
0.0014 ± 0.0010
0.0057 ± 0.0009
0.0091 ± 0.0029
0.0014 ± 0.0003
0.0019 ± 0.0009
0.0040 ± 0.0022
0.0014 ± 0.0004
0.0070 ± 0.0013
0.0986 ± 0.019
-19
n.d.
n.d.
+13
+15
-24
+15
-6
-31
+60
+32
-51
-21
-9
-18
+36
-2
Notes: Values in the tables are the mean value ±1 standard deviation from four replicates.
The last column reports the relative percent difference between the measured and expected
value. The expected value is the concentration calculated from the PAH concentration
obtained by summing the soil and biochar contributions (Table 3.1.1). This is expressed as
a relative percentage of (measured - expected)/expected × 100.
46
Obviously, the effect of biochar addition in soils on the level of PAHs will depend on
the background level of PAHs in the soil before treatment (Nam et al., 2009; Wilcke et al.,
2000), the concentration of PAHs in the original biochar, and the quantity of added
biochar. Then, environmental processes (evaporation, biodegradation, or abiotic
degradation) will affect the fate and levels of PAHs in amended soil. Due to the lipophilic
nature of the PAHs, these compounds tend to bioaccumulate in plants (Duxbury et al.,
1997; Parrish et al., 2006). Leafy vegetables typically accumulate higher levels of PAHs
from the soil system than companion fruit or root crops (Lei et al., 2011). The levels of
PAH observed in some of the biochars (see below) do posses levels that could be of
potential health and environmental concern, depending on the application rate, original soil
concentrations, and end-use for the soil.
3.1.3.4. Determination of EPA and EU PAHs in different biochar samples
The method developed in this study was applied to the determination of USEPA and
EU PAHs in a suite of ten biochar investigated in a previous study (Fabbri et al., 2012).
With the exception of biochar S-18 and S-19 (distillers grain) and S-17 (Macadamia nut
shells), all the other biochars were derived from woody biomass (Table 3.1.5). Almost all
16 USEPA PAHs were detected and quantified in the biochars, as well as several EU
PAHs. However, HMW EU PAHs were not detected (Table 3.1.5). The recovery of spiked
deuterated PAHs ranged between 60 and 100% and for all the samples an average of 78%,
78 and 75% for d-acenaphthene, d-phenanthrene and d-chrysene, respectively, with ∼10%
RSD each. Despite the difference in feedstock and process treatment the PAH levels were
quite similar (1–19 µg g-1). One sample (biochar S-17) was characterized by high levels of
PAHs. However, the literature reports examples of biochar with much higher
concentrations, some comparable to those observed on soot (Hilber et al., 2012;
Schimmelpfennig and Glaser., 2012). A large number of biochars investigated by Hale et
al. (2012) exhibited total PAHs in the 0.07–3.27 µg g-1 interval when produced from slow
pyrolysis from different biomass at temperatures between 250 and 900 °C, and higher
values (45 µg g-1) from gasification. These examples underline the variety of PAH levels
that could find in biochars. With few exceptions (S-17), naphthalene was the most
abundant PAH, in accordance to previous studies (Hale et al., 2012; Hilber et al., 2012;
Schimmelpfennig and Glaser, 2012; Freddo et al., 2012; Kloss et al., 2012), followed by
47
phenanthrene. However, it is interesting to note that benzo[a]pyrene was detected in all
biochars analyzed here, with concentrations ranging from 0.01 to 0.67 µg g-1.
Sample S-2 was biochar obtained from the fast pyrolysis of hardwood sawdust at 500
°C, while S-3 the same biochar stored 1 year in an open drum subject to environmental
conditions (Fabbri et al., 2012). Table 3.1.5 shows that the levels of LMW PAHs did not
change markedly, confirming the strong sorption of PAHs to biochar. However, Hale et al.
(2012) reported that artificial aging in aqueous solutions generally increased the
concentration of PAHs on biochar, probably due to the leaching of hydrophilic components
leaving the more hydrophobic biochar fraction.
Biochars S-18 and S-19 produced from the same feedstock (distiller grains) at similar
pyrolysis temperatures (350 and 400°C, respectively) exhibited significantly different PAH
concentrations (total USEPA 5.0 and 2.2 µg g-1) suggesting the importance of pyrolysis
conditions, as well as the role of temperature. A general trend has been observed of
increasing PAH contents at shorter pyrolysis times and high pyrolysis temperatures (Hale
et al., 2012). A detailed study on the presence PAHs in biochar samples produced from
woody and herbaceous biomass pyrolyzed at different temperatures showed that the
concentration of pyrogenic PAHs peaked at 500 °C, a common temperature in slow
pyrolysis (Keiluweit et al., 2012). Chagger et al. (2000) demonstrated through modelling
that PAHs are preferentially formed in a fluidized bed reactor versus a kiln style reactor,
due to unstable combustion reactions present in a fluidized bed reactor. Schimmelpfennig
and Glaser have underlined the importance of the particular technological process on the
sorbed PAH concentrations, with wood gasifiers associated with the highest levels of
PAHs on the solid residuals (Schimmelpfennig and Glaser, 2012). These authors proposed
the naphthalene/phenanthrene ratio and the total PAHs concentrations as factors to
differentiate pyrolysis processes between biochars. These hypotheses are also supported by
our data, since biochars that are created by slow pyrolysis at longer residency times in kiln
style reactors possess lower sorbed amounts of PAHs compounds.
48
Table 3.1.5. Concentrations of the 16 USEPA PAHs and (#) 15 EUPAHs (µg g-1 mean of two duplicates). (RB reference biochar; characteristics of
biochars from S-2 to S-20 were published elsewhere [Fabbri et al., 2012].)
Sample Id.
Nr.
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
PAHs
Naphthalene
Acenaphtylene
Acenaphthene
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
Cyclopenta[c,d]pyrene#
Chrysene#
Benzo[a]anthracene#
5-methylchrysene#
Benzo[b]fluoranthene#
Benzo[k]fluoranthene#
Benzo[j]fluoranthene#
Benzo[a]pyrene#
Indeno[1,2,3-cd]pyrene#
Dibenzo[a,h]anthracene#
Benzo[ghi]perylene#
Dibenzo[a,e]pyrene#
Dibenzo[a,h]pyrene#
Dibenzo[a,i]pyrene#
Dibenzo[a,l]pyrene#
Σ 16 EPA PAHs
#
Σ 15 EU PAHs
RB
S-2
S-3
S-4
S-5
S-15
S-16
S-17
S-18
S-19
S-20
1.75
0.03
0.03
0.07
0.71
0.13
0.3
0.35
0.001
0.09
0.09
0.01
0.13
0.1
n.d.
0.19
0.15
0.06
0.15
n.d.
n.d.
n.d.
n.d.
4.3
0.97
1.57
0.50
0.62
0.25
0.25
0.03
0.14
0.07
0.01
0.05
0.04
0.11
0.02
0.02
n.d.
0.02
0.02
0.02
0.01
n.d.
n.d.
n.d.
n.d.
3.6
0.32
1.71
0.30
0.31
0.16
0.30
0.04
0.08
0.07
0.01
0.02
0.02
0.04
0.05
0.01
n.d.
0.02
0.01
0.01
0.01
n.d.
n.d.
n.d.
n.d.
3.1
0.2
2.39
0.04
0.05
0.10
0.56
0.07
0.11
0.08
0.01
0.02
0.05
0.02
0.04
0.04
n.d.
0.10
0.13
0.01
0.01
n.d.
n.d.
n.d.
n.d.
3.8
0.43
0.44
0.01
0.01
0.05
0.31
0.03
0.08
0.08
0.01
0.02
0.04
0.02
0.04
0.02
n.d.
0.01
n.d.
0.01
0.01
n.d.
n.d.
n.d.
n.d.
1.2
0.18
0.47
0.02
0.07
0.08
0.27
0.03
0.11
0.12
0.01
0.03
0.04
0.02
0.05
0.02
n.d.
0.05
0.02
0.01
0.02
n.d.
n.d.
n.d.
n.d.
1.4
0.27
0.93
0.12
0.08
0.04
0.36
0.04
0.05
0.04
0.01
0.02
0.02
0.09
0.02
0.01
n.d.
0.02
0.01
0.01
0.01
n.d.
n.d.
n.d.
n.d.
1.8
0.22
2.58
0.71
0.28
0.92
3.88
0.65
2.46
2.58
0.05
0.92
0.83
0.27
0.70
0.43
n.d.
0.67
0.50
0.08
0.53
n.d.
n.d.
n.d.
n.d.
19
5.0
0.78
0.10
0.24
0.59
0.49
0.19
0.10
0.16
0.04
0.42
0.46
0.21
0.29
0.39
n.d.
0.32
0.27
0.21
n.d.
n.d.
n.d.
n.d.
n.d.
5.0
2.6
0.49
0.05
0.22
0.26
0.33
0.12
0.09
0.07
n.d.
0.17
0.08
n.d.
0.05
0.07
n.d.
0.06
n.d.
0.19
n.d.
n.d.
n.d.
n.d.
n.d.
2.2
0.62
3.36
0.10
0.11
1.13
2.70
0.33
0.21
0.10
0.03
0.09
0.17
0.21
0.07
0.06
n.d.
0.22
0.03
0.06
0.08
n.d.
n.d.
n.d.
n.d.
8.8
1.0
49
Given the values of total PAHs reported in Table 3.1.5, as well as those reported in the
literature (Hale et al., 2012; Freddo et al., 2012) for the slow pyrolysis biochars and the
level of biochar applications recommended in agriculture practices, the increased levels of
elevated PAHs in biochar amended soil is not of universal concern. However, as also seen
in our data and those from other studies (Keiluweit et al., 2012; Kloss et al., 2012), some
biochars do have levels of sorbed PAHs that do exceed existing and proposed guidelines
for the usage of specific materials (e.g. sludge, wood ash) on land (Keiluweit et al., 2012;
Freddo et al., 2012; Kloss et al., 2012) including commercial biochar (Hilber et al., 2012).
In addition, the bioaccumulation of PAH compounds in produce grown in biochar
amended soils requires further investigation. Therefore, the development of valid analytical
procedures for the determination of PAHs in biochar and biochar amended soils is critical.
3.1.3.5. Determination of EPA-PAH in EU-COST biochar samples
The method described in the previous sections was applied to three biochar samples
that were part of a laboratory exercise organised within the EU COST Action TD1107
(http://cost.european-biochar.org/en). The Working Group 1.
The three biochar samples BC1, BC2 and BC3 were produced with a PYREG® 500 –
III pyrolysis unit (PYREG GmbH, Dörth, Germany) from different feedstock (woodchip
sievings, paper sludge - wheat husks blend, sewage sludge) at similar conditions.
Information on the process is available in Bucheli et al. (2014).
The method was the same as described in the previous sections, however, two sets of
analyses were run under slightly different conditions. In the first set of samples (A) about
20 g were dried at 40 °C for 48 hours and the test-sample spiked with the surrogate PAH
mix (Supelco for EPA 525 containing acenaphthene-d10, phenanthrene-d10 and chrysened12; in a second set of samples (B) about 200 g were dried at 40 °C for 96 hours and
spiked with 16 EPA PAHs (prepared from Dr. Ehrenstorfer PAH-Mix 9 deuterated, 10 ng
µl-1).
The results are reported in Tables 3.1.6, 3.1.7 and 3.1.8.
Data obtained from the few laboratories participating to the PAH determination in the
ring trial are being evaluated by the WP1. Preliminary results would indicate that no major
deviations in the reported data occurred for the total PAH concentrations. Naphthalene was
the most dominant PAH in all the three biochars followed by phenanthrene. Naphthalene
was the PAH with the largest deviations, probably because of its volatility. The longer
50
drying period in test B seems to have resulted in loss of naphthalene.
Table 3.1.6. Concentrations of PAHs in biochar 1 in ng g-1.
BC1
A
mean
Naphthalene
Acenaphthylene
Acenaphthene
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
Chrysene
Benzo[a]anthracene
Benzo[b]fluoranthene
Benzo[k]fluoranthene
Benzo[a]pyrene
Indeno[1,2,3-cd]pyrene
Dibenzo[a,h]anthracene
Benzo[ghi]perylene
PAH TOTAL
1475
86
24
27
143
30
46
20
30
81
41
51
n.d.
n.d.
n.d.
2055
sd
56
11.4
4.1
7.1
49
8
11.7
6.6
5
7.6
5.5
12.5
-
51
B
mean
507
49
23
41
136
23
33
31
18
26
51
27
32
997
sd
84
6
5
12
41
3
7
4
6
3
10
3
1
Table 3.1.7. Concentrations of PAHs in biochar 2 in ng g-1.
BC2
A
mean
Naphthalene
Acenaphthylene
Acenaphthene
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
Chrysene
Benzo[a]anthracene
Benzo[b]fluoranthene
Benzo[k]fluoranthene
Benzo[a]pyrene
Indeno[1,2,3-cd]pyrene
Dibenzo[a,h]anthracene
Benzo[ghi]perylene
PAH TOTAL
1864
149
18
26
168
8
53
63
18
22
29
11
5.8
6.9
2.3
8.5
2454
sd
174
23
5
5.7
19
1.6
1.8
3.4
1.2
4.3
6.9
3.5
1.6
0.9
0.8
2.5
52
B
mean
1039
24
21
14
117
33
41
49
21
23
17
15
12
11
0
12
1450
sd
90
4
3
3
27
7
9
15
6
3
4
3
4
3
6
Table 3.1.8. Concentrations of PAHs in biochar 3 in ng g-1.
BC3
A
mean
Naphthalene
Acenaphthylene
Acenaphthene
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
Chrysene
Benzo[a]anthracene
Benzo[b]fluoranthene
Benzo[k]fluoranthene
Benzo[a]pyrene
Indeno[1,2,3-cd]pyrene
Dibenzo[a,h]anthracene
Benzo[ghi]perylene
PAH TOTAL
363
4.5
10.6
10.5
50.2
13.2
16.6
24.8
7.1
18.0
6.2
5.2
8.4
5.4
4.2
9.5
558
sd
83
1.1
0.8
1.6
5.8
1.5
1.5
2.4
1.1
3.4
1.5
1.3
2.3
1.6
0.8
3
B
mean
330
13
8
13
99
28
38
52
15
31
10
12
21
17
0
13
699
sd
33
4
0
3
13
2
5
6
2
5
2
1
3
2
1
3.1.4. Conclusions
A method for the determination of PAHs in biochar was developed making use of a
solvent mixture (1:1 acetone:cyclohexane) in place of more toxic and/or hazardous
solvents (e.g., dichloromethane, toluene) which was appropriate for the determination of
LMW PAHs (including naphthalene) along with HMW PAHs. The method was validated
with a certified reference soil and demonstrated its validity for the detection of PAHs
deriving from biochar in a soil matrix amended with 1% biochar. Because of the strong
affinity of PAHs toward biochar, solvent and duration time of the Soxhlet extraction were
crucial parameters and at least 36 h was necessary to obtain a satisfactory recovery with
1:1 acetone:cyclohexane. Furthermore, this method provided satisfactory recovery when
applied to a wide range of biochar samples obtained at different pyrolysis conditions from
different biomass parent materials suggesting that this analytical procedure could be used
successfully on different biochars. All the biochar analyzed contained the USEPA, as well
53
as some of the EU PAHs at detectable levels ranging from 1.2 to 19 µg g-1. In particular,
the presence of EU PAHs on biochar could be of concern when biochars with elevated
levels of PAHs are used in human food production due to the potential of contamination.
However, this aspect requires further investigations.
54
3.2. Quantitative determination of PAHs in an agricultural soil treated
with biochar
3.2.1. Introduction
Biochar application to soils has recently emerged as a potential strategy to sequester
carbon into agricultural soils, improve physical, chemical and biological properties of soil
and produce renewable energy (Glaser et al., 2002; Lehmann et al., 2006; Steiner et al.,
2007; Sohi et al., 2010; Uchimiya et al., 2010; Galinato et al., 2011; Vaccari et al., 2011;
Ventura et al., 2013). Biochar is known to improve soil water-holding capacity (Case et al.,
2012; Basso et al., 2013; Baronti at al., 2014) and plant water availability (Baronti et al.,
2014), decrease nutrients leaching (Güereña et al., 2013) and bioavailability of heavy
metals (Park et al., 2011), improve soil structure (Case et al., 2012) and stimulate soil
microbial activity (Kolb et al., 2009; Rutigliano et al., 2014), and finally, increase the pH
of soil, due to which it can be used on acidic soils with poor cultivation properties (Glaser
et al., 2002; Slavich et al., 2013).
In spite of the unquestionable advantages, there are also certain threats related with the
production and subsequent utilisation of biochar. Among threats, most frequently
mentioned is the contamination of biochar with polycyclic aromatic hydrocarbons (PAHs)
and heavy metals. While in the case of heavy metals their levels are at relatively low
values (Freddo et al., 2012) and depend on the content of trace metals in the initial
material, studies concerning PAHs indicate (Freddo et al., 2012; Hale et al., 2012; Hilber et
al., 2012; Keiluweit et al., 2012; Oleszczuk et al., 2013; Fabbri et al. 2013) that biochars
may be contaminated with those compounds to a significant degree. PAHs are formed
during the pyrolysis of organic matter (including biomass). PAHs are well known
carcinogenic and persistent pollutants and 16 PAHs are classified as priority pollutants and
are heavily regulated by the US EPA due to their carcinogenic, mutagenic or teratogenic
properties (Keiluweit et al., 2012). Therefore, the application of biochar to agricultural soil
could carry a significant risk to human health by contaminating soils. Moreover, PAHs in
soil may exhibit a toxic activity towards different plants, microorganisms and invertebrates
(Guo et al., 2012).
Several studies have shown that the presence of biochar in soils can influence the
bioavailability and bioaccessibility of organic contaminants (Beesley et al. 2010; GomezEyles et al. 2011; Zhang et al. 2013; Lattao et al. 2014). The biochar is particularly
55
effective at adsorbing and sequestering organic contaminants; enhanced sorption of
hydrophobic organic compounds such as PAHs could actually decrease microbial
mineralization by decreasing bioavailability (Rhodes et al., 2008, 2010; Xia et al., 2010),
and could lead to localize PAH accumulation. On the contrary, Liang et al. (2015) reported
that biochar amendment stimulates PAH-metabolizing bacterial activity by enhancing the
number of gene copies related to PAH degradation and by changing the structure of soil
microbial community.
Quilliam et al. (2012) reported that biochar can reduce the degradation of PAHs in two
important types of agricultural soil, which in the short term could not only increase the
concentration of soil PAHs but could also affect the long-term persistence of PAHs in the
environment. However, the environmental fates of biochar-associated PAHs added to soil
are still poorly understood. Therefore, it is necessary to improve understanding of the role
biochar plays in sorbing PAHs and on microbial activity and how this influences the
concentration of PAHs in soil and their persistence in the environment.
The aim of this study was quantifying the concentration of the 16 priority PAHs in soil
amended with biochar at two different rates and determine the impacts of biochar additions
on PAHs concentration in soils. Furthermore, the effect of biochar on PAHs levels was
compared with some properties of soil, which may influence the fate of PAHs.
3.2.2. Materials and Methods
3.2.2.1. Reagents and standards
PAH-Mix solution containing each of the 16 EPA PAHs [i.e., naphthalene,
acenaphthylene, acenaphthene, fluorene, phenanthrene, anthracene, fluoranthene, pyrene,
chrysene,
benzo[a]anthracene,
benzo[b]fluoranthene,
benzo[k]fluoranthene,
benzo[a]pyrene, indeno[1,2,3-cd]pyrene, dibenz[a,h]anthracene and benzo[ghi]perylene]
certified at concentrations of 10 mg l-1 for each species in acetonitrile was purchased from
Sulpeco (Belleforte, PA, USA).
Cyclohexane, acetone, ethyl acetate (all supra solv quality), and surrogate standard mix
(for EPA 525) containing acenaphthene-d10, phenanthrene-d10 and chrysene-d12 at
concentrations of 500 mg l-1 each in acetone were purchased from Sigma-Aldrich.
A solution of 1,3,5-tri-tert-butylbenzene (TTB, 10 mg l-1) in acetone:cyclohexane (1:1,
v/v) was prepared by weighing the pure compound purchased from Sigma-Aldrich.
56
3.2.2.2. Experimental layout
The field experiment was setup in a vineyard at the “Marchesi Antinori - La Braccesca
Estate” (Lat. 43° 10’ 15'’ N; Long. 11° 57’ 43’’ E; 290 m a.s.l.), located few kilometers
away from Montepulciano (Tuscany, central-Italy). The vineyard has been planted in 1995
and trellis system is a single curtain with plant-row spacing of 0.8 and 2.5 m; rows
orientation is East-West. The vineyard is not irrigated.
A randomized plot experiment, with three treatments and five replicates was setup in
2009. Each plot, 15 in total, had a surface area of 225 m2 (7.5 m in width and 30 m in
length) including 4 vineyard rows and 3 inter-rows. The treatments were: a single biochar
application at a rate of 22 t ha-1 in 2009 (B); two biochar applications at a rate of 22 t ha -1
each, in 2009 and 2010 (BB); and a control (C). Biochar was applied with two treatments,
in five replicates randomly distributed, as follows: 22 t ha-1 of biochar applied in 2009 (B);
22 t ha-1 in 2009 and further 22 t ha-1 in 2010 (BB) and control untreated plots (C).
Rows orientation is East-West, inter-rows are partially covered with spontaneous grass,
and tilled with a chisel plow in the March–June period. The vineyard is not irrigated and it
is fertilized with an inorganic fertilizer (15.0.26) twice per year (in November and April) at
a rate of 120 kg ha-1. Soil is acid, shallow and sandy-clay-loam textured (USDA, 2005)
(Table 1) and is highly compacted below 0.4 m depth
Untreated soil (control) and soil treated by two concentrations of biochar amendment
were sampled four times from 2011 to 2013 (August 2011, December 2011, May 2012 and
May 2013). Sixty samples (5 replicates x 3 treatments x 4 sampling seasons) were
examined, each sample was dried at 40 °C, sieved (mesh size: 2 mm) in order to obtain
homogeneous samples free of stones, larger roots and other coarse fragments, and stored at
- 20 °C. No losses of PAHs occur under these conditions.
3.2.2.3. Soil properties
Soils are shallow, acid, sandy-clay-loam (USDA, 2005) textured (Table 3.2.1). The
total organic Carbon content (C) and Total Nitrogen content (N) were analyze by dry
combustion elemental analyzer (Thermo Fisher Science) after fine grinding with a ball mill
to 0.5 mm. The pH was measured potentiometrically in a 1:2.5 soil– water suspension. The
CEC analysis was performed by saturation with barium–chloride at pH 8.2, displacement
57
of adsorbed barium by 0.05 M MgSO4 and titration of the Mg remaining in solution with
0.025M EDTA (Gessa and Ciavatta, 2000).
3.2.2.4. Biochar characterization
The biochar used in the experiment is a commercial charcoal provided by “Romagna
Carbone s.n.c.” (Italy) obtained from orchard pruning biomass through a slow pyrolisis
process at temperature of 500 °C in a transportable ring kiln of 2.2 m in diameter and
holding around 2 t of feedstock. The biochar at the end of the pyrolisis was crushed into
particles smaller than 5 cm of diameter before the soil application. Elemental composition
(C,H,N,S) was determined by combustion using a Thermo Scientific FLASH 2000 Series
CHNS/O Elemental Analyzer. Ash content of the biochar was measured by heating
samples in a muffle at 550 °C for 6 hours, as proposed by ANPA (2001). The oxygen
content was calculated from mass balance: %O=100-% (C+H+N+ash).
As the carbonate concentration of the soils was negligible, the total measured C
concentration was considered to represent total organic carbon (TOC). The TOC content
was determined on 5 samples of biochar amended soil and 5 control soil according to the
Ministero per le Politiche Agricole (1999), Method VII.1. Samples were pre-treated with
HCl 1.5 M (40 uL in 2-3 g of sample), heated at 60 °C for 1 hour; this procedure was
repeated for 4-5 times, till the samples stop reacting with HCl. Determinations were made
using a Thermo Scientific FLASH 2000 Series CHNS/O Elementar Analyzer.
A mixture of biochar with deionized water at 1:10 wt/wt ratio was prepared, thoroughly
mixed and pH measured at room temperature with a digital pH meter (HI 98103,
Checker®, Hanna Instruments). Prior to this analyses, biochar was sieved at 2 mm and
oven dried at 40 °C for 72 h.
3.2.2.5. Determination of polycyclic aromatic hydrocarbons
3.2.2.5.1. Extraction and clean up
The PAHs determined comprised of 16 compounds (US EPA List; Table 3.2.2).
Analyses of PAHs were conducted as described in Fabbri et al. (2013). Briefly, about 5 g
of sample was placed into the extraction cellulose thimble, spiked with 0.1 mL of surrogate
PAH mix (Supelco for EPA 525 containing acenaphthene-d10, phenanthrene-d10 and
58
chrysene-d12 5 μg mL-1 each in acetonitrile). The thimble was covered with cotton wool
and inserted into the Soxhlet extractor. The extraction was performed with 160 ml of
extraction solvents (acetone/cyclohexane (1:1, v/v)) mixture for 36 h (4 cycles h-1). The
extraction solvent was filtered, added with 1 ml of n-nonane, and then carefully evaporated
by rotatory vacuum evaporation at 40 °C.
The concentrated extract was collected and loaded onto a silica gel cartridge (6 ml, 1 g
DSC-Si Supelco washed with ethyl acetate, dried and conditioned with 4 ml cyclohexane).
After purification with 1 mL of cyclohexane, PAHs were eluted with 4 ml of
acetone/cyclohexane (1:1, v/v). The obtained solution was then blown down to 10 – 50 μl
under nitrogen, spiked with 10 μl of the internal standard solution 1,3,5-tri-tertbutylbenzene (TTB at 10 mg l-1) prior to GC–MS analysis.
3.2.2.5.2. GC-MS
Samples (1 µl) were injected under a splitless condition (1 min, then split ratio 1:50 to
the end of analysis) into a 6850 Agilent HP gas chromatograph connected to a 5975
Agilent HP quadrupole mass spectrometer. Analytes were separated by a HP-5MS fused
silica capillary column (stationary phase poly[5% diphenyl/95% dimethyl]siloxane, 30 m×
0.25 mm i.d., 0.25 mm film thickness) with the following temperature program: 50 °C to
100 °C at 20 °C min−1, then from 100 °C to 300 °C at 5 °C min−1, then a hold for 2.5 min
at 300 °C, using helium as the carrier gas. The mass spectrometer operated under electron
ionization (70 eV) and acquisition was performed on single ion monitoring (SIM) at the
molecular ion of each PAH at the time windows corresponding to the elution region of the
target PAH.
Acenaphthene-d10 was utilised to quantify naphthalene, acenaphthylene, acenaphthene
and fluorene; phenanthrene-d10 to quantify phenanthrene, anthracene, fluoranthene and
pyrene; chrysene-d12 to quantify the remaining PAHs. Recovery of surrogated PAHs was
determined with respect to the internal standard TTB. The procedural blank was
determined by going through the same extraction and cleanup procedures for each series of
samples. None of the analytical blanks were found to have detectable contamination of the
monitoring PAHs and thus the results were not blank corrected.
59
3.2.2.6. Statistical analysis
Mean and standard deviation of five replicates were used to compare results of soils
and biochar amended soils. An analysis of variance (ANOVA) test was conducted with R
software version 3.1.2 (2014-10-31) to evaluate significant difference between control and
biochar amended soil.
3.2.3. Results and discussion
3.2.3.1. Soil and biochar characteristics
The study was carried out using an agricultural soil classified as sandy-clay-loam
(USDA, 2005) textured with 70% sand, 15% silt and 15% clay. The soil characteristics
were as follows: pH 5.37, total C 0.77%, total N 0.24%, total H 0.43%, and cation
exchange capacity of 12.1 meq 100 g-1.
Results of biochar characterizations are reported in Table 3.2.1. The biochar used for
soil amendment had a total content of C, N, H, and O of 71.4%, 0.7%, 1.5%, 5.9%,
respectively, an ash content of 19.9% and a pH of 9.8 (Table 3.2.1). The biochar had a
molar H/C ratio of 0.26 and molar O/C ratio of 0.06, indicating a comparably high
aromaticity of the biochar carbon (Zimmerman et al., 2013).
Table 3.2.1. Chemical characteristics of biochar applied in the field experiment.
C (%)
Value
71.4
H (%)
1.54
N (%)
0.72
S (%)
0.59
O (%)
5.9
H/C (molar)
0.26
O/C (molar)
0.06
Ash (%)
19.9
pH
9.8
Charred (%)
97.6
60
The concentration of ∑16PAHs in the utilised biochar is 3.5 μg g-1 and all the US EPA
PAHs were detected, with naphthalene as the most abundant species followed by
phenanthrene (Table 3.2.2). With this concentration would pass current quality standards
by the European Biochar Certificate (4 μg g-1 for premium quality and 12 μg g-1 for basic
quality) and the International Biochar Initiative (6 μg g-1). Additional details about the
physicochemical properties of the biochar are presented in Baronti et al. (2014).
Table 3.2.2. BIOCHAR. Concentrations of the 16 USEPA PAHs and standard deviation
(n=2) in biochar applied in the field experiment.
Sample Id.
PAHs
BIOCHAR
ng g-1 SD
Naphthalene
2149
658
Acenaphthylene
42
12.0
Acenaphthene
37
2.1
Fluorene
60
6.7
Phenanthrene
674
38
Anthracene
92
1.4
Fluoranthene
133
0.2
Pyrene
150
33
Chrysene
49
11
Benzo[a]anthracene
109
18
Benzo[b]fluoranthene
98
3.6
Benzo[k]fluoranthene
88
34
Benzo[a]pyrene
93
6.9
Indeno[1,2,3-cd]pyrene
19
5.6
Dibenzo[a,h]anthracene
21
2.2
18
3834
2.1
819
Benzo[ghi]perylene
Σ 16 EPA PAHs
3.2.3.3. Total PAH in soil and in biochar amended soil
Almost all 16 US-EPA PAHs were detected and quantified in the amended soil samples
analysed (Table 3.2.3 and Table 3.2.4). In all of the untreated soils, acenaphthylene,
acenaphthene, indeno[1,2,3-cd]pyrene, dibenzo[a,h]anthracene and benzo[ghi]perylene
61
were not detected. The recovery of spiked deuterated PAHs ranged between 60 and 100%
and for all the samples an average of 78%, 78 and 75% for acenaphthene-d10,
phenanthrene-d10 and chrysene-d12, respectively, with ∼10% RSD each.
Total PAHs concentrations in untreated soils ranged from 18 ng1 to 29 ng g-1, in
amended soils B from 26 ng g-1 to 60 ng g-1and in amended soils BB from 192 ng g-1 to 60
ng g-1 (Table 3.2.3 and Table 3.2.4). The total level of 16 PAHs in amended soils BB was
on the same level as other data reported for agriculture soils in Europe, for example, 60145 ng g-1 for arable soil in Switzerland (Bucheli et al., 2004); 187 ng g-1 for rural soil in
United Kingdom (Wild and Jones, 1995) and 150 ng g-1 in Norwegian agriculture soil
(Nam et al., 2008). The levels in amended soils BB, instead, were significantly lower than
those in soils from non-industrial areas in China 318 ng g-1 and in arable lands in Poland
395 ng g-1 (Maliszewska-Kordybach et al., 2008).
Fig. 3.2.1. Concentrations of the 16 USEPA PAHs and standard deviation (n=5
plot) in soil, biochar amended soil BB and B at different sampling dates from the
beginning of the field experiment (May 2009).
200
180
Biochar amended soil BB
160
Biochar amended soil B
140
Soil C
ng g-1
120
100
80
60
40
20
0
Aug 2011
Dec 2011
May 2012
May 2013
After almost one year following biochar application, the total PAHs concentrations in
amended soils B resulted higher than those of the untreated soils, both in August (56 vs. 24
62
ng g-1) and December (39 vs. 24 ng g-1) (Fig. 3.2.1, Table 3.2.2 and Table 3.2.3).
Moreover, the differences were statistically significant although the high dispersion of
PAH values between samples withdrawn from the same parcel (n = 5) (Fig. 3.2.1). The
lower concentrations observed in winter for the treated soils suggest a seasonal variability
superimposed to sampling heterogeneity.
The mean concentration of total PAHs in soil amended BB, one year after the
application in August 2011, was approximately 6 times higher than the control soil (153 vs.
24 ng) (Fig. 3.2.1). However, the range of total PAHs concentrations in the 5 amended
soils BB sampled in August 2011 (102 ng g-1-192 ng g-1) remained within the range
reported for Italian agriculture soils (80-304 ng g-1, ARPA Piemonte - Rapporto Stato
Ambiente 2009).
Four years after the addition of 44 t ha-1 of biochar to agricultural soils BB, the PAH
concentration was significantly higher than that in unamended soil suggesting that biochar
can act as a source of soil contaminants. However, the level of PAHs in the biochar
amended soil soil remained within the maximum acceptable concentration for a number of
European countries, 5–50 µg g-1 (Carlon, 2007). In addition, table 3.2.5 shows that in the
amended soils BB the concentrations of the PAHs decreased significantly during the four
years following biochar application.
Quilliam et al. (2013) reported that biochar can reduce the degradation of PAHs in two
important types of agricultural soil, which in the short term could not only increase the
concentration of soil PAHs but could also affect the long-term persistence of PAHs in the
environment. On the contrary, our results suggest that the soil contamination by PAHs
following biochar application is not significant at the application rates currently
recommended in agriculture (20-60 t ha-1) and that biochar does not decrease PAH
degradation and has not long-term effects. Long-term impacts of biochar additions to soils
are still not fully understood, although evidence suggests that the characteristics of soil and
biochar are of central importance.
63
Table 3.2.3. Concentrations of the 16 USEPA PAHs and standard deviation (n=5
plot) in biochar amended soil B at different sampling dates from the beginning of
the field experiment (May 2009).
Sample Id.
PAHs
Aug 2011
ng g-1
Dec 2011
May 2012
May 2013
SD
ng g-1
ng g-1
ng g-1
SD
SD
SD
Naphthalene
16.9
5.5
9.9
0.9
7.2
2.2
6.6
1.9
Acenaphthylene
2.34
1.59
0.52
0.66
n.d.
-
n.d.
-
Acenaphthene
0.32
0.48
0.73
0.54
n.d.
-
n.d.
-
3.5
1.72
2.8
1.78
2.8
1.26
2.6
1.13
Phenanthrene
12.1
3.6
8.4
2.0
7.5
0.7
6.5
0.8
Anthracene
2.19
0.70
1.71
0.57
1.50
0.52
1.32
0.52
Fluoranthene
5.27
2.42
4.47
1.50
3.41
0.28
2.91
0.37
Pyrene
4.74
2.06
3.66
1.29
2.76
0.30
2.64
0.20
Chrysene
1.47
0.44
1.29
0.34
0.94
0.24
0.89
0.15
Benzo[a]anthracene
1.72
0.68
1.32
0.29
1.17
0.11
1.20
0.20
Benzo[b]fluoranthene
1.69
0.51
1.53
0.42
1.25
0.25
1.48
0.22
Benzo[k]fluoranthene
1.36
0.94
1.33
0.94
0.99
0.72
1.00
0.61
Benzo[a]pyrene
0.97
0.40
0.85
0.35
0.76
0.36
0.78
0.35
Indeno[1,2,3-cd]pyrene
0.69
0.44
0.50
0.37
0.18
0.39
0.30
0.52
Dibenzo[a,h]anthracene
n.d.
-
n.d.
-
n.d.
-
n.d.
-
Benzo[ghi]perylene
0.51
55.8
0.51
17.3
0.28
39.0
0.28
7.6
n.d.
30.5
4.5
n.d.
28.1
2.9
Fluorene
Σ 16 EPA PAHs
64
Table 3.2.4. Concentrations of the 16 USEPA PAHs and standard deviation (n=5
plot) in biochar amended soil BB at different sampling dates from the beginning of
the field experiment (May 2009).
Sample Id.
PAHs
Dec 2011
May 2012
May 2013
ng g-1
SD
ng g-1
ng g-1
ng g-1
Naphthalene
74.1
20.9
41.1
15.5
34.5
8.1
35.0
5.0
Acenaphthylene
1.7
0.3
1.1
0.4
1.0
0.4
1.0
0.5
Acenaphthene
2.0
1.0
1.4
0.5
1.5
0.4
1.1
0.2
Fluorene
4.8
1.4
2.7
0.6
2.4
0.6
2.4
0.5
29.8
11.5
17.5
5.9
18.7
5.4
14.3
5.5
3.5
0.6
2.2
0.9
2.2
1.0
1.5
0.4
Fluoranthene
10.3
3.3
5.9
2.3
6.8
3.5
4.9
1.7
Pyrene
10.3
3.5
5.8
2.2
6.7
3.4
4.9
1.7
Chrysene
3.0
1.0
2.0
0.6
1.4
0.8
1.2
0.3
Benzo[a]anthracene
2.9
0.8
2.2
0.9
2.3
0.9
1.9
0.6
Benzo[b]fluoranthene
3.8
1.8
3.3
1.8
2.9
1.1
2.8
1.1
Benzo[k]fluoranthene
2.5
1.6
1.8
0.9
1.6
0.9
1.6
0.9
Benzo[a]pyrene
2.3
0.6
2.0
0.4
2.2
0.6
1.8
0.3
Indeno[1,2,3-cd]pyrene
0.91
0.38
0.82
0.28
0.85
0.21
0.90
0.30
Dibenzo[a,h]anthracene
n.d.
-
n.d.
-
n.d.
-
n.d.
-
1.0
153.0
0.42
37.7
0.83
97.7
0.37
35.5
0.95
88.5
0.53
29.7
0.85
78.42
0.42
20.6
Phenanthrene
Anthracene
Benzo[ghi]perylene
Σ 16 EPA PAHs
Aug 2011
65
SD
SD
SD
Table 3.2.5. Concentrations of the 16 USEPA PAHs and standard deviation (n=5
plot) in soil control at different sampling dates from the beginning of the field
experiment (May 2009).
Sample Id.
PAHs
ng g-1
SD
ng g-1
SD
ng g-1
SD
ng g-1
SD
Naphthalene
5.6
2.0
6.2
1.3
5.0
2.0
5.6
1.0
Acenaphthylene
n.d.
-
n.d.
-
n.d.
-
n.d.
-
Acenaphthene
n.d.
-
n.d.
-
n.d.
-
n.d.
-
Fluorene
1.55
0.3
1.61
0.42
1.74
0.14
1.52
0.27
7.8
1.4
7.7
1.2
5.66
0.7
6.7
2.0
Anthracene
0.90
0.08
0.82
0.32
0.90
0.32
0.92
0.48
Fluoranthene
2.70
1.0
2.62
0.47
2.12
0.71
1.67
0.53
Pyrene
1.83
0.89
1.81
0.20
1.82
0.57
1.37
0.31
Chrysene
0.50
0.12
0.53
0.08
0.56
0.16
0.62
0.20
Benzo[a]anthracene
0.98
0.25
0.84
0.16
0.80
0.13
0.73
0.16
Benzo[b]fluoranthene
0.89
0.14
0.82
0.28
1.24
0.46
1.05
0.40
Benzo[k]fluoranthene
0.74
0.16
0.79
0.14
0.59
0.10
0.55
0.10
Benzo[a]pyrene
0.75
0.03
0.79
0.16
0.63
0.16
0.80
0.18
Indeno[1,2,3-cd]pyrene
n.d.
-
n.d.
-
n.d.
-
n.d.
-
Dibenzo[a,h]anthracene
n.d.
-
n.d.
-
n.d.
-
n.d.
-
Benzo[ghi]perylene
n.d.
24.2
3.08
n.d.
24.5
2.01
n.d.
21.0
3.97
n.d.
21.6
3.01
Phenanthrene
Σ 16 EPA PAHs
Aug 2011
Dec 2011
May 2012
May 2013
3.2.3.4. Individual PAH concentration and degradation
The individual concentrations of 16 US EPA PAHs in control soils and in soils
amended by two concentrations of biochar are presented in Tables 3.2.3, 3.2.4 and 3.2.5.
The PAHs with 2–3 rings composed the majority of PAHs in control soil and in amended
soil samples while PAHs with 4–6 rings only accounted for 24-40% of ∑PAHs on average.
A detailed analysis of the contribution of the individual PAHs in amended soils
indicated the dominance of naphthalene (40 ± 3.5% BB and 26 ± 3.2% B of the total
PAHs) and phenanthrene (20 ± 1.3% BB and 25 ± 1.5% B of the total PAHs) in all the
samples studied.
66
In the control soils, phenanthrene, naphthalene and fluoranthene dominated the PAH
profiles, supplying 30 ± 2.4%, 25 ± 1.3%, and 10 ± 1.5% of the total PAH concentrations,
respectively. In this control soil, the observed fingerprints of PAHs are in agreement with
data in the literature, where the same compounds were reported to be the dominating ones
in soil samples (Bucheli et al., 2004; Zhang et al., 2006; Kwon et al., 2014). Table 3.2.6
and 3.2.7 show that in the amended soils B and BB the concentrations of the PAHs
decreased significantly during the four years following biochar application. The total PAH
decreases by about 50% compared to that found the first year. However, the degradation
was generally higher for low molecular weight PAHs. Up to 61 and 56% in soils B and
BB, respectively, whereas for PAHs with five and six rings the corresponding figures
varied between 1 and 37%. In particular, naphthalene reduces its first year value to 61% in
B and 53 in BB. Acenaphtylene, acenaphthene, fluorene, phenanthrene, anthracene,
fluoranthene, pyrene and chrysene were all reduced to approximately 40-50% of their
initial values (concentration found the first year). The PAHs with higher molecular
weights,
benzo[a]anthracene,
benzo[b]fluoranthene,
benzo[k]fluoranthene,
benzo[a]pyrene, indeno[1,2,3-c,d]pyrene and benzo[ghi]perylene showed a decrease
between 1 and 37%.
In a few cases, an increase in concentration of PAHs is observed, which is likely caused
by the heterogeneity of the soil. High molecular weight PAHs are well known for their
recalcitrance to biodegradation because of their low bioavailability. PAH molecule
stability and hydrophobicity are two primary factors which contribute to the persistence of
HMW PAHs in the environment. This could due to the higher biodegradation rate of soil
bacteria in utilizing lower than higher ring PAHs as energy (Olson et al., 2008). However,
it is interesting to note that benzo[a]pyrene, considered to be representative for the group
of cancerogenic PAHs, was detected in all amended soil analyzed here, with concentrations
from 0.8 to 2.3 ng g-1. This levels of benzo[a]pyrene are lower than 3-13 ng g
-1
reported
for agriculture soils in Norway, Poland, Czech Republic and China (Nam et al., 2008;
Gusev et al., 2008; Maliszewska-Kordybach et al., 2009; Cao et al., 2013). Moreover, the
level of benzo[a]pyrene varied in untreated soils from 3 to 3.7%, in amended soils B and
BB from 1.5 to 2.8%. The presented results for control soil are in agreement with other
data reported in the literature (3-5%) concerning soils from non-industrial areas (Desaules
et al., 2008; Maliszewska-Kordybach et al., 2009; Cao et al., 2013).
67
Table 3.2.6. Reduction PAH concentration (%) in amended soils B compared to
concentration found the first year (August 2011) at different sampling dates from
the beginning of the biochar amendment (April 2009).
Dec
2011
27
moths
May
2012
37
moths
May
2013
49
moths
Naphthalene
41
57
61
Acenaphthylene
78
-
-
-128
-
-
Fluorene
21
21
26
Phenanthrene
31
38
46
Anthracene
22
31
40
Fluoranthene
15
35
45
Pyrene
23
42
44
Chrysene
12
36
40
Benzo[a]anthracene
23
32
30
Benzo[b]fluoranthene
10
26
13
Benzo[k]fluoranthene
2
27
26
Benzo[a]pyrene
12
22
19
Indeno[1,2,3-cd]pyrene
28
74
57
Dibenzo[a,h]anthracene
-
-
-
45
30
45
50
Sample Id.
PAHs
Time after
amendment
Reduction
(%)
Acenaphthene
Benzo[ghi]perylene
Total PAHs
68
Table 3.2.7. Reduction PAH concentration (%) in amended soils BB compared to
concentration found the first year (August 2011) at different sampling dates from
the beginning of the biochar amendment (April 2009).
Dec
2011
27
moths
May
2012
37
moths
May
2013
49
moths
Naphthalene
45
53
53
Acenaphthylene
33
40
41
Acenaphthene
32
26
43
Fluorene
44
51
51
Phenanthrene
41
37
52
Anthracene
38
38
56
Fluoranthene
42
34
52
Pyrene
44
35
52
Chrysene
33
52
61
Benzo[a]anthracene
26
21
37
Benzo[b]fluoranthene
14
25
28
Benzo[k]fluoranthene
28
35
37
Benzo[a]pyrene
11
5
19
Indeno[1,2,3-cd]pyrene
10
6
1
Dibenzo[a,h]anthracene
-
-
-
36
42
49
Sample Id.
PAHs
Time after
amendment
Reduction
(%)
Benzo[ghi]perylene
Total PAHs
69
Table 3.2.8. Reduction PAH concentration (%) in soils C compared to
concentration found the first year (August 2011) at different sampling dates from
the beginning of the biochar amendment (April 2009).
Dec
2011
27
moths
May
2012
37
moths
May
2013
49
moths
-1
11
0
Acenaphthylene
-
-
-
Acenaphthene
-
-
-
Fluorene
-3
-12
2
Phenanthrene
2
27
13
Anthracene
9
-1
-3
Fluoranthene
3
22
38
Pyrene
1
0
25
Chrysene
-7
-12
-25
Benzo[a]anthracene
14
18
25
Benzo[b]fluoranthene
8
-39
-18
Benzo[k]fluoranthene
-7
20
25
Benzo[a]pyrene
-5
17
-7
Indeno[1,2,3-cd]pyrene
-
-
-
Dibenzo[a,h]anthracene
-
-
-
-1
13
11
Sample Id.
PAHs
Time after
amendment
Reduction
(%)
Naphthalene
Benzo[ghi]perylene
Total PAHs
3.2.2.5. Molecular diagnostic ratios
PAH diagnostic ratios have been used to determine the source of PAH and the relative
importance of combustion and petroleum derived PAH in sediments and in soils (Yunker
et al., 2002; Tobiszewski and Namieśnik, 2012; Vane et al., 2013). PAH diagnostic ratios
may be an efficient supporting tool in studying the fate of PAH in the soil and assessing
the influence of the biochar on the PAH degradation/leaching. Mutsazawa et al. (2001)
investigated the photodegradation of PAHs emitted with diesel particles deposited on soils
and found that fluoranthene and pyrene were rather stable, but that pyrene degraded faster
70
on most of the model soils. Under natural conditions, the photodegradation of PAHs bound
to diesel particles deposited on soils is expected to be very slow.
The literature provides descriptions of more than ten different molecular diagnostic ratios
(Katsoyiannis et al. 2011; Tobiszewski and Namieśnik 2012). In this study, three
molecular
ratios
were
used:
anthracene/(phenanthrene+anthracene),
fluoranthene/(fluoranthene+pyrene),
naphthalene/(naphthalene+phenanthrene).
Anthracene/(phenanthrene+anthracene) and fluoranthene/(fluoranthene+pyrene) diagnostic
ratios
are
frequently
applied
to
soil
samples;
on
the
contrary
naphthalene/(naphthalene+phenanthrene) is not used in literature. Schimmelpfennig and
Glaser (2012) have found that naphthalene and phenanthrene can be specific of the source
of different biochars.
The results of calculations of molecular diagnostic ratios for the soil, biochar amended soil
and biochar samples are presented in Table 3.2.9 and in Fig. 3.2.2 in the form of the socalled
cross
plots.
The
values
of
anthracene/(phenanthrene+anthracene)
and
fluoranthene/(fluoranthene+pyrene) in the biochar amended soil B and BB were of 0.100.17 and 0.50-0.55, respectively, characteristic for contaminants of pyrogenic origin, and
thus biochar. Moreover, those results are in agreement with the results of Kuśmierz et al.
(2014) for soils situated in the vicinity of biochar production sites (Table 3.2.9). In
addition, are similarly with molecular ratios calculated for various biochars on the basis of
literature
data
(Table
3.2.9).
PAH
cross
plots
for
the
ratios
of
fluoranthene/(fluoranthene+pyrene) vs. naphthalene/(naphthalene+phenanthrene) shows
that biochar addition to the soil induced a decrease of these molecular diagnostic ratios
(Fig. 3.2.2). In particular, the biochar amendment at 44 t ha−1 caused a considerable
similarity of these diagnostic ratios to those of the used biochar.
PAHs present in soil can be degraded by native bacteria and fungi (Zhang et al., 2006),
resulting in a (possibly selective) decrease of concentration over time, with rates depending
on soil type, organic carbon and nutrient content, humidity and aeration (Sabaté et al.,
2006; Zhang et al., 2006). The results of microbial PAH degradation studies indicate that
phenanthrene may be degraded faster than anthracene, and fluoranthene faster than pyrene
(Sabaté et al., 2006). Moreover, PAHs may undergo desorption: fluoranthene and pyrene
are desorbed at similar rates, but phenanthrene is desorbed faster than anthracene (Enell et
al., 2005). The change of molecular ratios as a function of time for the biochar amended
soils B and BB are presented in Table 3.2.10. It is interesting to note that PAHs molecular
ratios have remained largely unchanged during the four years following biochar
71
application. Therefore, these resultants may indicate that PAH degradation in biochar
amended soil is influenced by biochar.
72
Table 3.2.9. Calculated anthracene/[phenanthrene+anthracene] (ANT/[ANT+PHE]), fluoranthene/[fluoranthene+pyrene] (FLA/[FLA+PYR]),
naphthalene/[naphthalene+phenanthrene] (NAP/[NAP+PHE]) ratios in biochar amended soil B and BB, soil C, biochar sample and literature
data.
ANT/[ANT+PHE]
FLA/[FLA+PYR]
NAP/[NAP+PHE]
Sample name
Present work
Soil C
0.1143
0.5683
0.4467
Soil B
0.1646
0.5382
0.5302
Soil BB
0.1038
0.5019
0.6934
Biochar
0.1201
0.4707
0.7611
Literature
Wood 400°C
0.1006
0.3715
-
Wood 500 °C
0.1238
0.3302
-
Wood 600 °C
0.0683
0.3667
-
Biochar 1
0.1923
0.4627
0.7714
Biochar 2
0.1571
0.5304
0.7330
Biochar 3
0.1668
0.5840
0.7876
Biochar 4
0.1705
0.5495
0.7621
Hardwood 500 °C
0.1071
0.6666
0.8626
Wood waste 550 °C
0.0882
0.5000
0.5866
Wood waste 470 °C
0.0879
0.5789
0.8101
Soil W1a
0.2396
0.4951
-
Soil W1b
0.1969
0.5303
-
Soil W1c
0.2139
0.4916
-
73
Keiluweit et al. (2012)
Hilber et al. (2012)
Fabbri et al. (2013)
Kuśmierz et al. (2014)
Table
3.2.10.
Calculated
anthracene/[phenanthrene+anthracene]
(ANT/[ANT+PHE]),
fluoranthene/[fluoranthene+pyrene]
(FLA/[FLA+PYR]), naphthalene/[naphthalene+phenanthrene] (NAP/[NAP+PHE]) ratios in biochar amended soil B and BB and soil C
sampled four times from 2011 to 2013 (August 2011, December 2011, May 2012 and May 2013).
ANT/[ANT+PHE]
FLA/[FLA+PYR]
NAP/[NAP+PHE]
Aug 2011
0.1032
0.5955
0.4186
Dec 2011
0.0964
0.5907
0.4475
May 2012
0.1378
0.5373
0.4679
May 2013
0.1199
0.5497
0.4528
Aug 2011
0.1532
0.5264
0.5833
Dec 2011
0.1696
0.5496
0.5424
May 2012
0.1668
0.5532
0.4912
May 2013
0.1688
0.5237
0.5038
Aug 2011
0.1044
0.4993
0.7130
Dec 2011
0.1099
0.5048
0.7019
May 2012
0.1041
0.5031
0.6482
May 2013
0.0969
0.5005
0.7104
Sample name
Soil C
Biochar amended soil B
Biochar amended soil BB
74
Fig. 3.2.3. PAH cross plots for the ratios of fluoranthene/[fluoranthene+pyrene] (FLA/[FLA+PYR]) vs.
naphthalene/[naphthalene+phenanthrene] (NAP/[NAP+PHE]).
0.60
0.58
0.56
FLA/[FLA+PYR]
0.54
0.52
Soil
0.50
Biochar amended soil B
0.48
Biochar amended soil BB
BIOCHAR
0.46
0.44
0.42
0.40
0.00
0.10
0.20
0.30
0.40
0.50
NAP/[NAP+PHE]
75
0.60
0.70
0.80
3.2.3.6. Effects of biochar on soil properties and relationship with PAHs
The chemical and physical characteristics of the soil and biochar amended soil B and
BB during the three years of studies are reported in Tables 3.2.11, 3.2.12 and 3.2.13.
Biochar amended soil pH, C content and CEC were increased and soil bulk density
decreased under biochar amendments in both treatments, being more or less in proportional
to the amendment rates for pH and CEC. On the contrary, the changes in C content and soil
bulk density in all years were not corresponding with the biochar amendment rates. In
particular, biochar amendment at 44 t ha−1 caused a consistent decrease in soil bulk density
by 0.15 mg m−3 in 2010 year and by 0.06 g cm−3 in 2011 and 2012 years as compared to
the corresponding control, respectively. Moreover, the biochar amendment at 44 t ha−1
caused a consistent increase in organic C by 4.7% in 2010 year, by 3.9% in 2011 year and
by 3.4% in 2012 year as compared to the corresponding control, respectively. This effect,
however, also if less consistent is present in the biochar amended soil B. In these soils the
biochar amendment at 22 t ha−1 caused an increase in organic C by 0.68% in 2010 year, by
0.49% in 2011 year and by 0.37% in 2012 year as compared to the corresponding control,
respectively.
Biochar addition to the soil induced a significant increase of soil CEC. This increase of
CEC in the biochar amended soils B and BB compared to control C (Table 3.2.11, 3.2.12
and 3.2.13) is proportional to the amendment rates. The increase of CEC in soils amended
can be attributed to the presence of retained oxygen content in biochar used in this
experiment, as previously published (Lee et al., 2010). The retained oxygen content
presents itself on the biochar as primarily carbonyl, carboxyl, and phenolic groups, all of
which in part facilitate CEC through electrostatic interactions. Furthermore, this data
shows that the biochar sample has the capability of not only serving as a long-term carbon
sequestration agent, but also has the potential to increase soil CEC.
The soil pH was clearly modified by the amendments in both treatments. In particular,
the pH of the soil the first year after application significantly increased with the
concentration of biochar amendment, starting from 5.50 in the control to 6.47 in the
biochar amended soil B and 7.18 in the BB (Table 3.2.11, 3.2.12 and 3.2.13). A similar
trend was observed by Fellet et al. (2011) using the same concentrations of biochar
application. Compared to the first year of biochar amendment, biochar amended soil pH
almost unchanged in the 2011 and 2012 in both treatments. The increase in soil pH after
the application of biochar may be attributed to the alkaline nature of biochar.
76
PAH degradation in the soil is slow; however, PAHs may be degraded through properly
stimulated soil microorganisms by mineralization, co-metabolic degradation and nonspecific radical oxidation (Wetzel et al., 1997). Soils inherently contain complex
autochthonous microbial communities, which have PAH degrading abilities (Ding et al.,
2010). The acidity of soils can control conditions for microbial degradation and regulate
the sorptive capacity of organic matter (e.g. by protonation of acidic functional organic
matter groups), thus it may contribute in different ways to soil matrix effects on PAH
degradation (Maliszewska-Kordybach, 1999; Bucheli et al., 2004). Therefore, in biochar
amended soil BB the higher pH may have been important for influencing the fate of PAHs.
Moreover, the biochar has been shown to increase microbial activity in soil (Steinbeiss et
al., 2009) which can stimulate PAH degradation. Our results showed that total PAH in BB
soil decreases by about 50% in two years, and therefore the addition of biochar could have
increased the degradation of PAHs.
Table 3.2.11. Chemical characteristics of soil C in the field experiment.
Soil
Jun 10
Feb 11
Jun 11
Jun 12
C (%)
0.67±0.02
0.73±0.08
0.82±0.07
0.93±0.04
pH
5.50±0.22
5.18±0.30
5.25±0.15
5.39±0.26
EC (meq 100 g-1)
12.6±0.4
11.8±0.9
11.5±1.5
11.9±1.9
Bulk density (mg m-3)
1.44±0.05
1.44±0.10
1.45±0.06
1.44±0.03
Notes: Values in the tables are the mean value ± standard deviation from five replicates.
77
Table 3.2.12. Chemical characteristics of biochar amended soil B in the field experiment.
Biochar amended soil
Jun 10
Feb 11
Jun 11
Jun 12
C (%)
1.35±0.25
1.33±0.30
1.31±0.29
1.30±0.29
pH
6.47±0.24
6.54±0.25
6.32±0.14
6.34±0.24
EC (meq 100 g-1)
18.16±0.96
18.32±1.05
18.14±0.83
18.22±0.78
Bulk density (mg m-3)
1.42±0.03
1.42±0.07
1.40±0.03
1.40±0.06
Notes: Values in the tables are the mean value ± standard deviation from five replicates.
Table 3.2.13. Chemical characteristics of biochar amended soil BB in the field experiment.
Biochar amended soil
Jun 10
Feb 11
Jun 11
Jun 12
5.4±1.2
5.11±0.96
4.76±0.53
4.3±1.4
pH
7.18±0.11
6.76±0.18
6.59±0.20
6.61±0.30
EC (meq 100 g-1)
26.0±3.7
24.3±1.8
24.1±1.8
22.9±1.8
Bulk density (mg m-3)
1.29±0.18
1.38±0.06
1.38±0.25
1.38±0.09
C (%)
Notes: Values in the tables are the mean value ± standard deviation from five replicates.
3.2.4. Conclusions
The fate of PAHs in biochar amended soil is relevant in order to prevent severe
contamination to the environment. The results presented in this study show that the biochar
addition determines an increase of the amounts of PAHs. However, the results
corresponding to the amendment of 22 and 44 t ha-1 suggest that the soil contamination by
PAHs following biochar application is not significant at the application rates currently
recommended in agriculture (20-60 t ha-1). In fact, the levels of PAHs in the soil remained
within the maximum acceptable concentration (5–50 µg g-1) for a number of European
78
countries. Moreover, biochar amendment in four years does not increase the concentration
of soil PAH. Therefore, the biochar does not reduce the degradation of PAHs in
agricultural soil and does not affect the persistence of PAHs in the environment. However,
the impact of biochar on the fate of PAHs needs to be investigated further for different
soils, biochars, over longer periods, and also under different field conditions.
79
4. Biochar characterization for agricultural utilization
4.1. Relationships between chemical characteristics and phytotoxicity of biochar from
poultry litter pyrolysis
4.1.1. Introduction
Biochar is the carbonaceous product of biomass pyrolysis which can be used as soil
additive capable to mitigate a variety of agro-environmental stresses through the
permanent storage of biomass carbon, pH correction, reduced synthetic fertilizer use,
decreased runoff of fertilizers and agrochemicals (Glaser et al., 2002; Lehmann et al.,
2006; Steiner et al., 2007; Sohi et al., 2010; Uchimiya et al., 2010; Galinato et al., 2011;
Vaccari et al., 2011; Ventura et al., 2013).
The effect of adding biochar to soils may result in increased plant growth, productivity
and yield (Graber et al., 2010, Vaccari et al., 2011, Joseph et al., 2013) attributed to the
improvement of soil water-holding capacity (Case et al., 2012; Basso et al., 2013; Baronti
at al., 2014), lower disease incidence in crops (Matsubara et al., 2002; Elad et al., 2010;
Elmer and Pignatello, 2011), reduced bioavailability of heavy metals (Park et al., 2011),
increased nitrogen and carbon bioavailability (Scharenbroch et al., 2013). Preventing loss
of nutrient leaching may reduce the needs of fertilizer use (Liang et al., 2006; Laird et al.,
2010). Because of its basicity, biochar can be used in acidic soils with poor cultivation
properties (Glaser et al., 2002; Slavich et al., 2013). However, the effects of biochar are
highly variable depending on the feedstock, thermochemical process conditions,
application rate, soil characteristics, environmental conditions, and plant species (Chan and
Xu, 2009; Jeffery et al., 2011, Schulz and Glaser, 2012), explaining the variety of
outcomes reported in literature that range from a boost in plant productivity to evident
phytotoxicity (Jeffery et al., 2011).
Prompted by the urgency to find applications alternative to its disposal and
management, poultry litter has been investigated as a substrate in the preparation of
biochar (Chan et al., 2008; Van Zwieten et al., 2013; Novak et al., 2014). Possible benefits
of amending soils with poultry litter biochar have been reported and attributed to an
improved nitrogen availability (Chan 2008; Van Zwieten et al., 2013). Lower N2O
emissions with respect to the raw poultry litter and the elimination of potential pathogens
has advocated its pyrolytic conversion (Van Zwieten et al., 2013). However, the use of
80
biochar especially from animal origin has raised concerns related to its possible toxicity
and studies have been recently conducted to explore physiological effects on biota (Bastos
et al., 2014; Smith et al., 2013; Oleszczuk et al., 2012).
Bioassays based on seed germination and early stage seedling growth is a simple and
commonly used ecotoxicological test for evaluating the impact of biochar amendment on
crop growth (Solaiman et al., 2012). The test of phytotoxicity of the biochars was made in
the absence of soil due to the large soil–char interactions observed in some studies
(Zimmerman et al., 2011) and because Solaiman et al. (2012) demonstrated that growing
seedlings in pure biochar materials is a valid tool in assessing the effect of biochar
application rate on germination.
Seed germination, one of the most important phases in the life cycle of a plant, is
highly responsive to existing environment (Kuriakose et al., 2008). Factors such as heavy
metals (Wollan et al., 1978), PAHs (Rogovska et al., 2012), ammonia (Wong et al., 1983),
salts (Adriano et al., 1973) and low molecular weight fatty acids (Zucconi et al., 1985)
have been shown to be responsible for inhibitory effects.
Some studies have examined the effect of biochar on seed germination (Free et al.,
2010; Solaiman et al., 2012; Busch et al., 2013). Rogovska et al. (2012) reported that
biochars contain phytotoxic compounds that inhibit germination of maize. In contrast, Free
et al. (2010) reported that maize seed germination was not significantly affected by
biochars made from a range of organic sources. Solaiman et al. (2012) showed that
biochars generally increased germination at low application rates (10-50 t ha-1), whereas
higher application rates of 100 t ha-1 had no effect or decreased germination rate.
Alburquerque et al. (2014) observed that different biochars exerted a positive effect on
seed germination also to high application rates instead.
Recent studies have also suggested different methods for reducing the toxicity of
biochar (Bargmann et al., 2013; Buss and Masek, 2014). Washing biochar with water or an
organic solvent has been successfully tested to reduce phytotoxicity of biochar (Bargmann
et al., 2013; Bernardo et al., 2010; Rogovska et al. 2012). Meanwhile, Kołtowski et al.
(2015) demonstrated significant reduction of biochars toxicity by drying them at various
temperatures (100–300 °C) for 24 h.
While the published and ongoing investigations are providing increasing data helpful to
understand the relationships between biochar characteristics and seed germination
(Bargmann et al., 2013; Kołtowski et al., 2015), further studies are needed to better clarify
the role played by the chemical properties in determining the plant toxicity in order to
81
forecast strategies in biochar synthesis or post-treatments. Biochars from different
feedstock and process conditions may exhibit a wide range of plant response, from growth
inhibition to stimulation. The relatively simple seed germination test is a valid and fast tool
to compare several biochars obtained from different starting materials and under different
pyrolysis conditions. Since the test is performed in short time and without the buffering
effect of soil, it could be considered a kind of precautionary procedure that highlights
intrinsic phytotoxicity of the tested materials.
The aim of this study was to evaluate phytotoxicity of biochar from poultry litter by
means of standard germination tests and to identify possible relationships with its chemical
characteristics. To this purpose, germination tests with cress (Lepidium sativum L.) were
conducted to poultry litter biochars synthesised at different pyrolysis conditions. Besides
manure, poultry litter typically contains bedding materials made up of lignocellulosic
residues. Therefore, a comparison was made with biochars from a representative
herbaceous residue, corn stalk, prepared under the same conditions. The effect of solvent
extraction and biological conditioning on seed germination was tested on a selected poultry
litter biochar prepared upon pyrolysis at 400 °C (PL400). The chemical composition of
mobile constituents in this sample capable to be potentially released in the water and air
compartments was investigated by solvent extraction and solid-phase microextraction
(SPME).
4.1.2. Materials and Methods
4.1.2.1. Biochar synthesis
Cornstalk was described in a previous study (Cordella et al., 2012). Granular poultry
litter was a marketed organic fertilizer obtained after processing raw poultry litter collected
from local broiler farms by pasteurizing at 80–110°C, milling, and pelletizing. Biomass
was air dried at 60°C, milled and sieved at 2 mm before pyrolysis.
Batch pyrolysis experiments were conducted under nitrogen with a fixed bed tubular
quartz reactor placed into a refractory furnace (see Conti et al., 2014 for details) with about
20 g cornstalk or 35 g poultry litter exactly weighed and uniformly placed onto a sliding
quartz boat; nitrogen flow was set at 1500 cm3 min-1 and when the temperature inside the
reactor, measured with a thermocouple, reached the selected value, the boat was pushed
into the oven and left for a given residence time before pulling it back into the unheated
82
part of the reactor. Pyrolysis were performed under three different conditions based on a
previous study (Conti et al. 2014) of temperature/residence time: 400°C/20 min, 500°C/10
min and 600°C/5 min. In accordance to the original biomass (cornstalk, CS; poultry litter,
PL) and pyrolysis temperature, the obtained biochar samples were named CS400, CS500,
CS600, PL400, PL500 and PL600. Chemicals were purchased by Sigma Aldrich. SPME
Carboxen-PDMS fibers and the fiber holder were purchased by Supelco.
4.1.2.2. Biochar Characterization
Elemental composition (HCNS) was determined by combustion using a Thermo
Scientific Flash 2000 series analyzer. Ash was determined as the residual mass left after
exposure at 600 °C for 5 hours. The oxygen content was calculated from the mass balance:
O%=100-(C+H+N+ash)%.
Analytical pyrolysis (Py-GC-MS) were conducted at 900 °C for 100 seconds with a
CDS 5250 pyroprobe interfaced to a Varian 3400 GC-Saturn 2000 MS. GC-MS conditions
and the determination of indicators of carbonisation % charred and toluene/naphthalene
ratio were described in Conti et al. (2014).
The content of the 16 EPA priority PAHs was measured in triplicate as described in
Fabbri et al., (2013). Briefly, about 0.5 g of biochar were spiked with 0.1 mL of surrogate
PAH mix (Supelco for EPA 525 containing acenaphthene-d10, phenanthrene-d10 and
chrysene-d12
5
μg
mL-1
each
in
acetonitrile)
and
soxhlet
extracted
with
acetone/cyclohexane (1:1, v/v) for 36 hours. The solution was filtered, added with 1 ml of
n-nonane (keeper), carefully evaporated by rotatory vacuum evaporation at 40 °C and
cleaned up by solid phase extraction onto a silica gel cartridge before analysis with a
Agilent HP 6850 GC coupled to a Agilent HP 5975 quadrupole mass spectrometer; GCMS conditions were those detailed in Fabbri et al., (2013). Recovery of surrogate PAHs
was determined with respect to the internal standard tri-tert-butylbenzene added prior to
GC-MS analysis.
Volatile fatty acids (VFAs) were determined by the single drop extraction procedure as
described in Torri et al., (2014). About 200 mg of biochar exactly weighed was added with
0.1 ml of internal standard solution (1.0 g l-1 2-ethylbutyrate in deionised water) and
thoroughly mixed with 0.2 ml of saturated aqueous KHSO4. After centrifugation, a drop of
dimethyl carbonate (1.2 µl) from a 10 µl chromatography microsyringe was exposed into
the supernatant aqueous solution. After 20 min exposure the drop was retracted and
83
injected into a GC-FID (injection temperature 250°C) equipped with polar GC column
(Agilent Q7221J&W nitroterephthalic-acid-modified polyethylene glycol DB-FFAP 222
30 m, 0.25 mm, 0.2 µm) with the following thermal program: 80°C for 5 min, then
10°C/min to 250°C. Calibration was performed by applying the same procedure to
standard solutions containing known concentration of each VFA (namely: acetic,
propionic, isobutyric, butyric, isovaleric and valeric acid).
For the determination of N-NH4+, about 10 g of biochar were placed in an end-to-end
shaker for 2 h with 1 N KCl (1:10 dw:v) followed by centrifuging at 4500 x g for 20 m and
passing through a 0.45 µm paper filter.
4.1.2.3. SPME of mobile compounds
The SPME was applied to aqueous extracts following the procedure under development
by Ghidotti et al. 2014 using a Carboxen-PDMS fiber direclty immersed (DI-SPME) into
the test solution added with KH2PO4/Na2HPO4 phosphate buffer 2M at pH 5.3 and internal
standard (o-eugenol and 2-ethyl butyric) under magnetic stirring for 30 minutes followed
by thermal desoprtion at 250 °C and GC-MS analysis.
Head space (HS) analysis was performed following the procedure described by Spokas
et al., 2011 modified for the SPME sampling with Carboxen-PDMS fiberv (HS-SPME)
utilising o-eugenol as internal standard (Ghidotti et al. 2015).
Separation of thermally desorbed compounds was conducted with a DB-FFAP polar
column (30m lenght, 0.25mm i.d, 0.25µm film thickness).
4.1.2.4. Biochar post-treatments. Aqueous extraction
About 2 g of PL400 was extracted with 50 mL of deionised water in a 100 mL flask at
room temperature for 12 hours with mechanical shaking. The aqueous phase was separated
by filtration through a 0.22 mm paper filter and used as such for the germination test, while
an aliquot was kept at –20 °C for SPME-GC-MS analysis (see above). The solid biochar
residue left after water extraction was further extracted with 50 mL of methanol under
reflux for 12 hours. The methanol was separated by filtration and an aliquot corresponding
to the 40g/L suspension of biochar was poured into petri dishes and dried overnight at 70
84
°C under vacuum to remove all the methanol. Thereafter deionised water was added to
perform germination tests. The final solid biochar residue left after water and methanol
extraction was dried overnight at 100°C under vacuum and utilized for germination tests.
4.1.2.5. Biochar post-treatments. Biological treatment
Microbial treatment of PL400 was conducted for 14 days with an activated sludge. The
sludge was obtained from a municipal wastewater treatment plant located in Ravenna after
centrifugation at 6000 rpm (20% w/w volatile suspended solids). A suspension containing
0.5 g of the concentrated sludge and 250 ml of 40 g l-1 PL400 in deionised water was
thoroughly mixed under laminar shake at 120 rmp overnight. An aliquot of 10 ml of this
suspension was added in petri dishes and stored at 14 days at 25°C before performing
germination test as shown above.
4.1.2.6. Germination tests
The germination tests were conducted in four replicates by incubating 50 seeds of cress
(Lepidium sativum L.) with 5 g of a mixture containing biochar and deionized water onto
sterilized cellulose filter paper (Whatman No. 1) placed in a petri dish sealed with paraffin
film. Three levels of biochar concentration were tested 2, 5 and 40 g L-1. These rates were
equivalent to 2, 5 and 40 t ha-1 on an area basis of 10 cm soil depth and a dry bulk density
of 1.5 kg m-3. Germination tests were also performed on the fractions obtained from the
chemical and biological post-treatments of PL400 described above. The quantities of these
fractions were adjusted to correspond to the concentration level of 40 g l-1 of the original
biochar. Before incubation, the samples were shaken at 150 r.p.m. on a platform shaker at
room temperature for 24 h. pH and electrical conductivity (EC) were determined. The pH
was directly measured placing the glass-electrode into the suspension with a pH-meter
Mettler Toledo SG 2-ELK. The electrical conductivity (EC) was measured with a Delta
OHM HD 8706 conductimeter in the supernatant obtained by centrifuging the suspension
and filtered at 0.45 micron.
Phytotoxicity tests were performed on biochar:deionized water mixtures (wetted
biochar) according to the procedure described in UNI 11357:2010. The experiments were
conducted with 50 seeds of cress which were incubated with 10 g of biochar saturated with
deionized water according to value of the water holding capacity (table 4.1.1) on sterilized
85
cellulose filter paper placed in a petri dish. All Petri dishes were covered and incubated in
room thermostat at 25 ± 2° C for 72 ± 0.5 hours in the dark. Similarly, a control was
prepared with deionised water.
After 72 h of exposure, a visible root development was used as the operational
definition of seed germination. Data were reported as percentage relative seed germination
(RSG) with respect to the control (deionised water):
RSG = (number of seeds germinated in the sample/ number of seeds germinated in
control) * 100
4.1.2.7. Statistical analysis
All the experiments were conducted at least in duplicate. Results of germination tests
were evaluated statistically using Analysis of Variance (ANOVA) performed with
STATISTICA (StatSoft Italia, 2011) and GMAV (Underwood and Chapman, 1997)
followed by Student-Newman-Keuls post hoc tests.
4.1.3. Results and Discussion
4.1.3.1. Characterization of biochar
Bulk analysis
The yields and characteristics of biochars obtained from the pyrolysis of poultry litter
(PL400, PL500, PL600) under three different pyrolysis conditions are reported in Table
4.1.1 and compared with those of biochar from corn stalk (CS400, CS500, CS600). As
expected, the chemical characteristics of the biochars were dependent on the original
biomass and the pyrolysis conditions. In particular, the H/C and O/C ratios decreased with
increasing pyrolysis temperature, while the content of ash increased as observed with the
same pyrolysis unit under the same conditions (Conti et al. 2014). Biochar from poultry
litter contained higher levels of nitrogen, sulphur and ash, derived from the manure
fraction, as demonstrated by comparative studies on manure and lignocellulosic biochars
(Novak et al., 2014). In general, the elemental composition, ash content and the trends with
pyrolysis conditions of the poultry litter chars here investigated were comparable to those
reported in the literature (Chan et al. 2008, Cimò et al., 2014, Song et al., 2012, Van
Zwieten et al., 2013).
86
Extractable compounds
The concentrations of specific potentially toxic extractable compounds, namely PAHs,
VFAs and ammonium are reported in Table 4.1.2. Solvent extractable PAHs occurred
within the range of 0.7 - 1.7 mg kg -1, values that were typical of biochars from different
origins (Hale et al., 2012; Fabbri et al., 2013; Hiber et al., 2012). PAHs concentration can
be considered negligible for acute effect lower than typical values in soils, Bucheli et al.,
2004. Generally, naphthalene was the most abundant PAHs followed by phenanthrene and
fluorene, the level of benzo[a]pyrene was in the 5-65 ng g-1 range.
The concentration of VFAs was significant higher in poultry litter biochars (4-9 mg kg1
) in comparison to those from cornstalk. (2-4 mg kg-1). Acetic acid was always the most
abundant VFA. VFAs derived from feedstock fermentation during silage in poultry litter
and the new formation of fatty acids from the thermal degradation of lipids.
Ammonium was not detected in cornstalk biohar, whereas it was abundant in biochars
from poultry litter with higher concentrations in the less carbonised biochars.
87
Table 4.1.1. Yields, water-holding capacity, elemental analysis and ash (% wt dry weight mean values ± s.d. n=4) and elemental molar ratios of
biochars from the pyrolysis of corn stalk (CS) and poultry litter (PL) at different conditions (400 °C/20 min, 500 °C/10 min, 600 °C/5 min).
waterBiochar
Yield
holding
(% wt)
capacity
(%)
Molar
Elemental content (%)
C
H
N
O
S
Ash
ratios
(%)
H/C
CS400
38.3±0.9
69.5
50±2.1
3.3±0.1
0.96±0.03
15±1.9
0.07±0.01
28.95±0.01
0.79
CS500
33±1.6
81.1
51±1.6
2.7±0.1
0.91±0.04
14.9±0.1
0.03±0.04
30.14±0.02
0.63
CS600
31.4±0.5
73.7
50.7±0.3
2.4±0.1
0.81±0.03
13±1.6
-
32.30±0.02
0.57
PL400
49±3.4
88.6
33±4.7
2.7±0.5
3.6±0.8
11±2.1
1.7±0.5
46.64±0.01
0.98
PL500
41.4±0.9
94.1
33±1.0
2.1±0.1
3.4±0.1
6.5±0.9
2.2±0.1
52.29±0.04
0.76
PL600
39.5±0.5
92.3
31.4±0.5
1.7±0.1
3.2±0.5
4.6±0.3
2.3±0.1
56.82±0.01
0.65
88
Table 4.1.2. Molecular analysis of extractable compounds and volatile matter by Py-GCMS of biochar from corn stalk (CS) and poultry litter (PS) (mean values and s.d. from two
replicates, T/N toluene/naphthalene ratio).
Extractable
Biochar
PAHs
VFAs
Py-GC-MS
NH4+
%
mg kg-1
T/N
mg g-1 mg kg-1 charred
CS400
0.72±0.06 3.8±1.2
-
53±3
8.5±0.7
CS500
1.09±0.05 0.9±0.3
-
80±3
5.4±1.5
CS600
0.84±0.01 2.6±0.1
-
92±3
3.0±1.7
9.3±0.3
45
88±9
13±10
PL500
0.88±0.05 4.3±2.0
25
90±3
12±6
PL600
0.79±0.01 6.8±1.8
14
88±3
11±2
PL400
1.7±0.2
Py-GC-MS. Thermolabile fraction
The molecular composition of the thermally labile fraction could be inferred from the
structural identification of the compounds identified in the pyrolysates (Conti et al., 2014,
Fabbri et al., 2013, Kaal and Rumpel, 2009). The pyrolysate of CS400 was characterised
by a complex pattern of compounds dominated by phenols and methoxyphenols associated
to the presence of partially charred lignin, while the pyrolysate of CS600 contained few
peaks due to the hydrocarbons associated to more heavily charred fraction (Fig. 4.1.1).
Proxies of the degree of carbonisation established in previous studies (Conti et al., 2014)
for lignocellulosic biomass were confirmed in this study for corn stalk biochar: the
toluene/naphthalene ratio and the relative abundance of compounds representative of the
charred fraction (% charred) exhibited a clear trend with H/C ratios (Table 4.1.2).
However, biochar samples from poultry litter did not exhibit significant changes with the
H/C ratios and the variability was higher. This finding would suggest that the pyrolysis
proxies developed for lignocellulosic biochar could not be valid for biochar containing
charred proteins and lipids.
The occurrence of partially charred components from proteins and lipids were clearly
evidenced in the pyrolysates of biochar. Phenols and methoxyphenols were detected in
PL400 as well as in CS400 in accordance to the fact that the original substrates contained a
89
Figure 4.1.1. Total ion chromatograms from Py-GC-MS of poultry litter (PL) and
cornstalk (CS) biochars.
90
lignocellulosic component. The distinctive signature of PL pyrolysate was the occurrence
of nitrogen-containing compounds (NCCs) from proteins and a pattern of n-alkanes/n-alk1-enes assigned to the thermal cracking of bound or free fatty acids. The occurrence of
saturated alkyl domains was confirmed by
13
C-NMR studies on poultry manure biochars
that disappear after carbonisation and may play a role in the availability of sorbed
compounds (Cimò et al., 2014). Among the tentatively identified NCCs, pyrrole, pyridine,
(iso)quinoline and carbazole along with their alkyl derivatives are indicative of partially
charred proteinaceous matter. It worthwhile to note that NCCs were also identified in the
volatile fraction by SPME as described in the next section.
SPME-GC-MS. Volatile and water soluble compounds
Information on the molecular characteristics of the mobile fraction was gathered by
HS-SPME (volatile) and DI-SPME (water soluble). The attention was specifically focused
to organic compounds, being reported that heavy metals are generally present below the
limits causing adverse effects and loosely bioavailable (Cely et al., 2015). Moreover,
Bastos et al. 2014 argued that in aqueous extracts PAHs and metals might occur at
concentrations below the level to pose detrimental effects, at least for woody biochar up to
80 t ha-1; however, it is to be remarked that the biological response depended on the
organisms selected in the bioassay (Bastos et al., 2014).
SPME was applied to sample PL400 that was utilised in post-treatment studies. The
results are shown in figure 2 for the analysis of volatile organic compounds (VOCs) by
HS-SPME (fig. 4.1.2A) and DI-SPME of aqueous extracts (fig. 4.1.2B), respectively. The
VOCs were characterised by the presence of a wide array of compounds deriving from the
thermal degradation of polysaccharides (e.g. cyclopentenones, furans), lignin (e.g. 4vinylphenol, guaiacol), NCCs (e.g. pyrroles, pyridines, indole), lipids (e.g. VFAs, acetic
acid is also derived from hemicellulose). Alkylated pyrazines and acetamide were probably
derived from Maillard reactions between carbohydrates and proteins.
Notably, a suite of short chain n-alkanes/alkenes was identified supporting Py-GC-MS
results and literature data about the occurrence of aliphatic components in poultry litter
biochar (Cimò et al., 2014). It is expected that the polar fraction of VOCs will be
preferentially distributed into the aqueous phase in comparison to non-polar constituents.
In fact, the SPME-GC-MS analysis of the PL400 water extract showed a predominance of
organic acids, including C2-C10 aliphatic and C7-C9 aromatic acids (Figure 4.1.2B).
91
Figure 4.1.2. Total ion chromatograms obtained after (A) HS-SPME of volatile organic
compounds and (B) DI-SPME of water extract of poultry litter biochar (PL400).
O
O
Counts
O
OH
OH
OH
OH
O
2.6e+07
B
MeO
OH
2.4e+07
i.s.
O
2.2e+07
O
OH
i.s.
2e+07
OH
O
1.8e+07
OH
O
1.6e+07
OH
O
1.4e+07
O
COOH
OH
1.2e+07
OH
O
1e+07
COOH
OH
8e+06
O
6e+06
OH
4e+06
2e+06
10.00
12.00
14.00
16.00
18.00
20.00
22.00
24.00
26.00
28.00 Minutes
OH
N
O
N
6e+07
i.s.
C5
A
C2
S
5.5e+07
O
O
O
N
S
5e+07
4.5e+07
MeO
CHO
O
Counts
OH
C2
O
O
C6-C10
hydrocarbons
N
N
4e+07
O
O
N
NH 2
N
3.5e+07
3e+07
OH
O
2.5e+07
O
OH
C3
OH
2e+07
1.5e+07
1e+07
5e+06
2
4
6
8
10
12
14
16
18
20
22
24
26
28
Minutes
4.1.3.2. Effect of biochar on seed germination
The pH and EC of the biochar/water suspensions utilized in germination tests are
reported in Table 4.1.3. The pH values were higher for the suspensions with the more
carbonized biochars from the same feedstock and increased for each biochar type with
increasing concentration. Under the same conditions, the pH was higher in the suspensions
with poultry litter biochar, in accordance to previous studies (Novak et al., 2014).
The EC values of the suspensions increased with increasing biochar concentration and,
at the same concentrations, the biochar from poultry litter had much higher (from 4 to 40
92
times for the same condition) EC values than the CS suspensions. Salinity can have a
detrimental effect on seed germination and plant growth, especially in the seedling stage,
though the response of various plant species to salinity differs considerably (Mengel and
Kirkby, 1987). In general, salinity effects are mostly negligible in extracts with EC
readings of 2000 µS cm-1 or less (Hoekstra et al. 2002). This critical level was exceeded in
poultry litter biochar at 40 g L -1. The toxicity of inorganic nitrogen results mainly from
ammonia (NH3) which affects plant growth and metabolism at low concentration levels at
which NH4+ is not harmful (Mengel and Kirkby, 1987). At concentrations of 0.15-0.20
mM, which are comparable to those calculated in the 40 g l-1 biochar/water mixtures, NH3
could be toxic (Bennett and Adams, 1970).
The effect of the biochar suspensions on seed germination of cress (Lepidium sativum
L.) is presented in Table 4.1.4 in terms of percent seed germination with respect to control
(deionised water only).
The assay results in this work suggested that all the cornstalk biochar suspensions had
little impact on seed germination as one-way ANOVA analysis showed no significant
difference between control group and test groups (p > 0.05). Noticeably, CS400 was
almost non-toxic to germination even if used as the growth substrate (UNI test, table
4.1.3).
On the contrary, all the biochar samples from poultry litter inhibited significantly the
seed germination at the highest level of 40 g L-1 in water suspensions. At the harsh
conditions of the UNI test the germination was totally suppressed.
The comparison with cornstalk suggested that the toxicity of biochar from poultry litter
could be explained by some distinctive chemical components originated from this
feedstock. Compounds derived from lignin and cellulose/hemicellulose could be excluded
on the ground that biochar samples from corn stalk did not suppress seed germination in
water suspensions. The suspensions of biochars from both substrates presented similar pH
values, thus this parameter is not involved in toxicity. This in accordance to the findings by
Gell et al. 2011 who did not evidenced clear trends of pH and short term phytotoxicity in
biochars of different origins, at least under neutral/basic conditions. Similarly, the
concentration of solvent extractable PAHs was similar in PL and CS biochars, thus PAHs
cannot be responsible of the observed toxicity. Acetic acid was present in all the biochars
and at similar levels, partly due to the decomposition of cellulose/hemicellulose. The suite
of alkanes/alkenes characterising the Py-GC-MS pyrolysates of poultry litter biochar
93
samples would suggest the presence of a lipid fraction producing shorter chain fatty acids
by thermal degradation as confirmed by SPME-GC-MS on PL400.
The main differences between the CS and PL biochars were the higher content of
elemental nitrogen (table 4.1.1) and ammonium (table 4.1.2), and the presence of a
thermally labile fraction derived from proteins and lipids (Py-GC-MS data).
Table 4.1.3. Results from chemical analysis of biochars and relative water suspensions.
Electrical conductivity
Biochar
(mS cm−1)
pH
2 g l-1 5 g l-1 40 g l-1
2 g l-1
5 g l-1 40 g l-1
CS400
7.6
8.0
8.5
16
75
1.8 103
CS500
8.2
8.5
9.3
72
76
1.9 103
CS600
8.4
8.9
10.1
1.9 102 3.5 102 1.9 103
PL400
8.0
9.0
9.7
7.1 102 1.3 103 7.3 103
PL500
8.4
9.3
10.2
8.7 102 1.5 103 7.7 103
PL600
9.4
9.8
10.3
9.3 102 2.0 103 8.1 103
4.1.3.3. Germination tests after biochar post-treatment
A selected sample of poultry litter biochar (PL400) was extracted with water followed
by methanol extraction, the extracts and the residue were utilized in germination tests.
Germination tests were also performed to PL400 after treatment with sewage sludge to
assess the effect of biodegradation. The quantity of extracts and the residues corresponded
to the biochar loading level of 40 g l-1. The results are presented in figure 4.1.3. The cress
germination rate in the water extracts was similar to that of the original biochar
suspensions indicating inhibition due to some components in the water extracts. The
germination rates increased significantly to values similar to the control when the
suspension was made with the biochar left after solvent extraction. These observations are
supported by the results of Rogovska et al. (2012), who showed that growth inhibition no
longer occurred when biochars were washed prior to germination.
94
Biochar suspensions treated with an active sludge for almost two weeks displayed a
germination rate similar to the extracted biochar (figure 4.1.3). The reduced toxicity could
be ascribed to microbial degradation of some noxious components as suggested by
Bargmann et al. (2013).
Table 4.1.4. Relative seed germination (% of control) of cress (Lepidium sativum L.) in
biochar:deionized water suspensions (2, 5, 40 g l-1) and phytoxicity tests (% seed
germination with respect to control) according to UNI 11357:2010 (mean values and s.d.
from four replicates). The percent seed germination in pure deionised water (control) is
also reported.
Germination (%)
Biochar
Id.
2 g l-1
5 g l-1
40 g l-1
UNI
Control
96±2
97±2
98±2
94±2
CS400
98±2
93±1
96±4
81±3
CS500
97±5
98±1
96±2
41±3
CS600
95±4
95±2
96±1
14±4
Control
92±2
92±2
92±2
94±1
PL400
83±4
74±3
53±19
no germination
PL500
77±2
73±2
47±10
no germination
PL600
77±3
74±3
53±7
no germination
4.1.4. Discussion
The results showed in Figure 4.1.3 indicated that the relative seed germination of water
extracts is low and comparable to that of the original biochar strongly supporting the
hypothesis that the polar/ionic constituents ending up in water are responsible to the
observed biochar toxicity. Similarly, Bargmann et al., 2013 applying germination tests to
hydrochars from various origins demonstrated that the inhibiting effects were caused by
some water soluble substances. These authors hypothesized that organic acids could be
possibly responsible of the toxicity of the water extractable fractions. The potential of
microbial detoxification was evidenced by Busch et al. (2013) who observed that the
genotoxicity of hydrochar mixed with compost became lower than that of pure hydrochar.
95
Figure 4.1.3. Seed germination rates relative to control of original poultry litter biochar
(PL400), water extracts and PL400 after post-treatments (solvent extraction and treatment
with active sludge). Gerimantion tests referred to a biochar load of 40 g l-1. The
germination rate of control (water only) is reported for comparison.
100
96
90
82
84
80
% germination
70
60
60
52
50
40
30
20
10
0
Biochar
Aqueous
Extract
Biochar after Biochar after
extraction biodegradation
Control
Gell et al., 2011 showed that the short term phytotoxicity of biochar is dependent on the
feedostock and is probably associated to ionic water soluble constituents rather than the
less polar organic compounds composing tars. In accordance, the methanolic extract of
biochar after water extraction exhibited a seed germination rate of 93% (not reported in
figure 4.1.3) higher than that measured in the water extracts. Interestingly, among the
various biochars investigated by Gell et al. (2011) those obtained from poultry biochar
exhibited positive effects (radish roosh elongation) and acted in decreasing phytotoxicity
of digestates. The calculated concentrations of VFAs in biochar suspensions (from data of
table 4.1.2) were higher than those that may cause detrimental effects, for instance
calculated PL400 VFAs at 40 g l-1 (374 µg g-1 ) was higher than 252 µg g-1 EC50 values
for plant growth (Himanen et al., 2012). Probably because different factors are governing
the physiological response of VFA including pH and bio-availability (Paavola and Rintala,
2008; Himanen et al., 2012). In addition, the occurrence of aromatic acids was identified
by DI-SPME along with VFAs.
96
These results of this study suggested that toxic compounds responsible for the toxicity
of PL400 were water extractable and biodegradable. The SPME analysis of the water
extracts (figure 4.1.3) evidenced that aliphatic and aromatic carboxylic acids were the
dominant compounds. Py-GC-MS and HS-SPME analyses evidenced the presence of
NNCs that seemingly were not partitioned into the water phase. The role of PAHs can be
excluded as they are not water soluble and occurred at low, and comparable to cornstalk,
levels in PL400.
Biochar has a potential as a soil amendment for improving soil quality, decrease
fertilizers losses and store carbon into the soil. Nevertheless, as soil additive, the absence
of phytotoxicity is the minimal requirement. Biochar from poultry litter may exert negative
effect at least at the relatively high level of soil amendment (40 t ha-1) due to the presence
of water soluble and biodegradable components, probably derived from the thermal
decomposition of proteins and lipids. However, the toxicity can drastically be reduced by
means of washing with water or mixing with biologically active material. Whereas
leaching (accompanied by wastewater generation) would be not an applicable option,
biological treatment (e.g. composting or mixing with activated sludge) of phytotoxic
biochars could be a simple and economic solution for increase the agronomic performance
of biochar characterized by toxicity issues. Results obtained shows that biochar are not an
“intrinsically safe” material, and every biochar (from different process and/or feedstock)
has to be evaluated, checked and eventually treated before the agronomic application.
97
5. Application of analytical pyrolysis methods to the characterization of
organic carbon in biochar
5.1. Characterisation of soil and biochar amended soil by hydropyrolysis
5.1.1. Introduction
Strategies to improve soil quality and increase the soil organic carbon (SOC) in
agricultural soils receive a lot of attention. SOC is an important soil constituent influencing
soil and water quality, farming practices and ultimately food production (Bruce et al.,
1998). Besides its significance to soil quality and food production, soil carbon pool plays
an important role in the overall global carbon budget.
A possible way to increase SOC content is to add biochar to soil (Lehmann et al., 2006;
Lehmann, 2007). Biochar is carbonaceous product of biomass pyrolysis which attracts
research interest due to its potential value for long-term carbon sequestration with
additional agronomic benefits. The application of biochar to soil has been proposed for
increasing the SOC and restraining the growth of atmospheric CO2 concentration
(Lehmann, 2007). Although our understanding of biochar stability has improved in recent
years (Ameloot et al., 2013), there is limited research on the effects of biochar on native
SOC and biochar carbon stability in soils in environmental conditions over a longer timescale. It is well known that a variable component of biochar is labile (degradable on
annual/decadal timescales) and hence, only a proportion of total carbon in biochar provides
long-term carbon sequestration.
Actually, an increasing number of observations suggests that biochar can be degraded,
by both biotic and abiotic processes (Hamer et al., 2004; Cheng et al., 2008; Guggenberger
et al., 2008). However, in most of the studies the stability of biochar was assessed during
laboratory incubations, with fresh biochars added to soil (Zavalloni et al., 2011; Ameloot et
al. 2013). The duration of these experiments ranges from several weeks (Cross and Sohi,
2011) to several years (Kuzyakov et al., 2009 and Kuzyakov et al., 2014), allowing to
understand biochar stability under controlled laboratory conditions. Moreover, in recent
studies various analytical techniques have been applied to investigate stability of biochar
(De la Rosa et al., 2008; Kaal et al., 2008, 2009; McBeath and Smernik, 2009; Michel et
al., 2009; Conti et al., 2014).
However, there are only few studies estimating biochar degradation rates in soil
(Kuzyakov et al., 2009; Hilscher and Knicker, 2011) and the long-term stability of biochar
98
in soils. This is because the changes of biochar content are too small for any practical
experimental period. Many studies estimating the decomposition rates of biochar in soil are
based on changes of CO2 efflux after biochar application. This approach is unsuitable to
estimate biochar decomposition because of the much higher contribution of soil organic
matter and plant residues mineralization of the CO2 compared to biochar.
Therefore, our study addresses the separation and determination of labile and resistant
carbon fractions in soils and biochar amended soils for the quantification of stabile fraction
(black carbon). An emerging pyrolytic approach isolating and quantifying BC in soils and
chars is hydropyrolysis combined with GC-MS. This analytical method has not yet been
applied to biochar amended soils in long term studies.
HyPy is pyrolysis assisted by high hydrogen pressures (150 bar) in presence of a
dispersed sulphided molybdenum catalyst. Application of HyPy to sediments, soils or
organic matter results in the reductive removal of all labile organic matter (defined as nonBCHyPy) (Wuster et al., 2012), so isolating a highly stable portion of the BC (BCHyPy) that is
predominantly composed of >7 ring aromatic domains (Meredith et al., 2012). The high
hydrogen pressure and slow heating rate employed, together with the presence of a
sulphided molybdenum catalyst, prevent the generation of secondary char (Love et al.,
1995) that is encountered with other chemical or thermal oxidative methods. In general,
HyPy offers a potential mean to discriminate between bound and adsorbed organic species.
As a result, the technique has been used to remove adsorbed products, facilitating analysis
of organic carbon in samples (Brocks et al., 2003).
It was also observed that HyPy appeared able to discriminate between relatively labile
biochars reporting low BCHyPy values and more refractory, high-BCHyPy soot in pure
samples, and between environmental samples from industrial sites with BC predominantly
derived from combustion of fossil fuels and agricultural sites dominated by the burning of
vegetation (Meredith et al., 2012).
The aim of the study consists of the identification of the BC and characterization of the
labile organic carbon in biochar amended soils in a four years field study. Here, for the first
time, we present the molecular composition of labile fraction of soil with biochar by HyPy.
99
5.1.2. Materials and methods
5.1.2.1. Soils collection and incubation experiment
The field experiment was setup in a vineyard at the “Marchesi Antinori - La Braccesca
Estate”, Montepulciano, Tuscany, Italy (43°10′15″ N, 11°57′43″ E). A randomized plot
experiment, with three treatments and five replicates, was setup in 2009. Each plot, 15 in
total, had a surface area of 225 m2 (7.5 m in width and 30 m in length) including 4
vineyard rows and 3 inter-rows. The treatments were: two biochar applications at a rate of
22 t ha-1 each, in 2009 and 2010 (BB); and a control (C). Biochar was applied in the interrow space of the vineyard using a spreader and it was incorporated into the soil using a
chisel plow tiller at 0.15 m depth.
Untreated soil (control) and soil treated by biochar amendment were sampled four
times from 2011 to 2013 (August 2011, December 2011, May 2012 and May 2013). Forty
samples (5 replicates x 2 treatments x 4 sampling seasons) were examined, each sample
was dried at 40 °C, sieved (mesh size: 2 mm) in order to obtain homogeneous samples free
of stones, larger roots and other coarse fragments, and stored at - 20 °C.
5.1.2.2. Soils, biochar amended soils and biochar characterization
The soil is a sandy-clay-loam (USDA, 2005) from the 0−30 cm horizon of the
vineyard. It was air-dried (72 h) and then sieved (2 mm). The contents of carbon, nitrogen,
hydrogen and sulfur of soil and biochar amended soil were determined by an elemental
analyzer (Thermo Scientific, FLASH 2000 Series). As the carbonate concentration of the
soils was negligible, the total measured C concentration was considered to represent TOC.
The pH, was measured potentiometrically in a 1:2.5 soil–water suspension. The CEC
analysis was performed by saturation with barium–chloride at pH 8.2, displacement of
adsorbed barium by 0.05 M MgSO4 and titration of the Mg remaining in solution with
0.025M EDTA (Gessa and Ciavatta, 2000). The texture of the vineyard soil was composed
of 15% clay, 15% silt, and 70% sand.
The biochar used in the experiment is a commercial charcoal provided by “Romagna
Carbone s.n.c.” (Italy) obtained from orchard pruning biomass through a slow pyrolisis
process at temperature of 500 °C in a transportable ring kiln of 2.2 m in diameter and
100
holding around 2 t of feedstock. The biochar at the end of the pyrolisis was crushed into
particles smaller than 5 cm of diameter before the soil application.
The contents of carbon, nitrogen, hydrogen and sulfur of biochar were determined by
combustion using a Thermo Scientific FLASH 2000 Series CHNS/O Elemental Analyzer.
Ash content of the biochar was measured by heating samples in a muffle at 550 °C for 6
hours, as proposed by ANPA (2001). The oxygen content was calculated from mass
balance: %O=100-% (C+H+N+ash). The pH of biochar was measured (1:10 wt/wt ratio of
biochar with deionized water) by a digital pH meter (HI 98103, Checker®, Hanna
Instruments) at room temperature. Prior to this analyses, biochar was sieved at 2 mm and
oven dried at 40 °C for 72 h.
5.1.2.3. Hydropyrolysis
Hydropyrolysis (HyPy) tests were performed using the procedure described in detail in
a number of publications (e.g. by Ascough et al., 2009; Meredith et al., 2012). Briefly, 50100 mg of biochar sample and 3-4 g of biochar amended soil were loaded with a Mo
catalyst using an aqueous/methanol 0.2 M solution of ammonium dioxydithiomolybdate
[(NH4)2MoO2S2]. Catalyst weight was ~ 5% of the sample weight for soil and biochar
amended soil, ~ 10% for biochar. The catalyst loaded biochar samples were placed within
shortened borosilicate pipette ends (20 mm long), plugged at each end with pre-cleaned
quartz wool and then placed in the HyPy reactor. The catalyst loaded soil and biochar
amended soil samples instead were placed directly in the reactor with steel wool on the
bottom. We used the recommended temperature program previously optimized for
pyrogenic carbon quantification where the samples are heated at rate of 300°C min-1 from
50 to 250°C, then heated at 8 °C min -1 from 250°C until the final temperature of 550°C for
2 min (Ascough et al., 2009; Meredith et al., 2012), all under a hydrogen pressure of 15
MPa. A hydrogen sweep gas flow of 5 L min-1, measured at ambient temperature and
pressure, ensured that the products were quickly removed from the reactor vessel, and
subsequently trapped in a silica gel-filled trap cooled by dry ice.
5.1.2.4 Black carbon quantification
The BC (reported as BCHyPy) content of each sample was derived by comparing the
organic carbon (OC) content of the catalyst loaded samples prior to HyPy with those of
101
their HyPy residues (Eq. (1)). Elemental composition (HCNS) was determined by
combustion using a Thermo Scientific Flash 2000 series analyzer.
BCHyPy (BC=OC%) =
Residual OC (mg C in HyPy residue including spent catalyst)
Initial OC (mg C in sample including catalyst)
x 100
As the carbonate concentration of the soils was negligible, the total measured C
concentration was considered to represent total organic carbon (TOC). The TOC content
was determined on 5 samples of biochar amended soil and 5 control soil according to the
Ministero per le Politiche Agricole (1999), Method VII.1. Samples were pre-treated with
HCl 1.5 M (40 uL in 2-3 g of sample), heated at 60 °C for 1 hour; this procedure was
repeated for 4-5 times, till the samples stop reacting with HCl. Determinations were made
using a Thermo Scientific FLASH 2000 Series CHNS/O Elementar Analyzer.
5.1.2.5. Non-BCHyPy fraction characterisation
The non-BCHyPy fraction (hydropyrolysate) from the soil, biochar amended soil and
biochar samples were desorbed from the silica recovered from the trap with 10 ml aliquots
of n-hexane and dichloromethane (DCM). The eluents were evaporated to 1 ml at room
temperature for 12 h prior to analysis. GC–MS analyses in full scan mode (m/z 35–650)
were performed on 6850 Agilent HP gas chromatograph connected to a 5975 Agilent HP
quadrupole mass spectrometer (EI mode, 70 eV), equipped with an autosampler and a
split/splitless injector. Analytes were separated by a HP-5MS fused silica capillary column
(stationary phase poly[5% diphenyl/95%dimethyl]siloxane, 30 m × 0.25 mm i.d., 0.25 mm
film thickness), using helium as the carrier gas, and an oven programme of 50°C (hold for
2 min) to 300°C (hold for 33 min) at 5°C min -1. Samples (1 µl) were injected under
splitless conditions (1 min, then split ratio 1:50 to the end of analysis) with an injector
temperature of 280°C. The abundance of the individual n-alkanes were quantified from the
m/z 57 mass chromatograms, and for the PAHs the mass chromatograms of the molecular
ion of each compound was used, following the addition of 100 µl of hexatriacontane (100
mg l-1, Sigma-Aldrich) and 100 µl of 1,3,5-tri-tert-butylbenzene (TTB, 100 mg l−1 SigmaAldrich) respectively as internal standards, assuming a response factor for each compound
of 1.
102
The PAHs were identified by matching the retention times of each peak in the sample
chromatogram with those of a standard solution. Interfering coelution problems were
evaluated in the samples by comparing mass spectra of the samples with those of the
standards as well as with those from the NIST mass spectra library (NIST MS Search r.
2.0).
5.1.2.6. Statistical analysis
Quantitative data are presented as mean values ± standard deviation (n = 5). An
analysis of variance (ANOVA) was undertaken to determine significant difference between
control and soil with biochar. A significant difference was statistically considered at level
of p < 0.05.
5.1.3. Results and discussions
5.1.3.1.
Properties of the soil and biochar
The study was carried out using an agricultural soil classified as sandy-clay-loam
(USDA, 2005) textured with 70% sand, 15% silt and 15% clay. The soil characteristics
were as follows: pH 5.37, total C 0.77%, total N 0.24%, total H 0.43%, and cation
exchange capacity of 12.1 meq 100 g-1
Results of biochar characterizations are reported in Table 5.1.1. The biochar used for
soil amendment had a total content of C, N, H, and O of 71.4%, 0.7%, 1.5%, 5.9%,
respectively, an ash content of 19.9% and a pH of 9.8 (Table 5.1.1). The biochar had a
molar H/C ratio of 0.26 and molar O/C ratio of 0.06, indicating a comparably high
aromaticity of the biochar carbon (Zimmerman et al., 2013). All the US EPA PAHs were
detected in the utilised biochar and summed up to 3.5 μg g-1, with naphthalene as the most
abundant species followed by phenanthrene. With this concentration would pass current
quality standards by the European Biochar Certificate (2013, 12 μg g-1) and the
International Biochar Initiative (2012, 20 μg g-1). Additional details about the
physicochemical properties of the biochar are presented in Baronti et al. (2014).
103
Table 5.1.1. Chemical characteristics of biochar applied in the field experiment.
Value
C (%)
71.4
H (%)
1.54
N (%)
0.72
S (%)
0.59
O (%)
5.9
H/C (molar)
0.26
O/C (molar)
0.06
Ash
19.9
pH
9.8
Charred (%)
97.6
5.1.3.2 Stable carbon fraction
It is important to determine how much of the carbon contained in biochar is potentially
stable over long periods of time as there are likely to be various fractions, differing in their
stability, ranging from very unstable (labile) fractions to very recalcitrant (stable) fractions.
The HyPy method has been demonstrated to remove almost all labile organic carbon,
leaving a residue of highly stable with the average ring structures greater than 7 fused rings
(Meredith et al., 2012; Wuster et al., 2012). The low molecular weight non-BCHyPy that are
removed by HyPy along with any other residual labile organic compounds are unlikely to
be stable on centennial timescales due to their susceptibility to biological and chemical
oxidation (Ascough et al., 2008).
We used the HyPy to quantify the effect of biochar addiction in soils on the level of
BC. Obviously, the effect of biochar addition in soils on the level of BC will depend on the
background level of BC in the soil before treatment, the BC in the original biochar, and the
quantity of added biochar. Then, environmental processes (evaporation, biodegradation, or
abiotic degradation) will affect the fate and levels of BC in amended soil.
The high BCHyPy value found for biochar sample (83±3.3%) and the very high stability
of this fraction under HyPy conditions (17±1.2%) seen in this study suggest that it is
104
composed predominantly of soot-derived BC and that carbon added with biochar should be
quite recalcitrant to degradation.
Table 5.1.2. Weight loss (%) of the soil and biochar amended soil during HyPy treatment.
Weight loss (%)
Sample
Aug 2011
Dec 2011
May 2012
May 2013
Biochar amended soil
6.2±0.7
6.9±0.4
5.8±0.8
7.7±1.2
Soil
7.0±0.8
7.1±0.5
7.4±0.3
6.6±0.4
Mass losses during HyPy of the samples were c. 7.7–5.8% w/w and no difference was
found observed between biochar amended soil and soil without biochar (Table 5.1.2). On
the contrary, the results show a significant difference in BCHyPy concentration during four
years of biochar experiment (Fig. 5.1.1). The BCHyPy concentration in amended soils, one
year after the application in August 2011, was approximately 20 times higher than the
control soil (79% ± 4.0% vs. 5.4% ± 1.3%). For the soil samples the BC HyPy content is
6.1% (±2.1%), which is comparable with range of BC contents reported by Hammes et al.
(2007) for a sand-rich soil (Chernozem) and a clay-rich soil (Vertisol). The carbon present
in control without biochar was very labile during HyPy, with complete conversion
apparent at 550°C, and therefore a BCHyPy content very low (BC/OC = 5.2-8.0%). The low
BCHyPy reported for soil samples suggests that is composed predominantly of
lignocellulosic material and humic acids.
105
BC as proportion of OC (%)
Figure 5.1.1. Black carbon (BC) as proportion of organic carbon (OC) as measured by
HyPy of the soil (C), biochar amended soil (BB) and biochar.
100.0
90.0
80.0
70.0
60.0
50.0
40.0
30.0
20.0
10.0
0.0
C
BB
Biochar
Biochar
Aug 2011
Dec 2011 May 2012 May 2013
The BCHyPy concentration in biochar amended soils in the 4-year study ranged between
79% ± 4.0% and 61% ± 2.4% (Fig. 5.1.1) The change BCHyPy as a function of time for the
biochar amended soils and reference soils without biochar are presented in Fig. 5.1.1. It is
interesting to note that the labile carbon during the first year was 21% and that really 21%
and 24% of total organic carbon has been lost after 2 e 3 years, respectively (Fig. 5.1.1).
However, table 5.1.2 shows that in biochar amended soil the BCHyPy decreased during the
three years following biochar application. The BCHyPy loss during the experiment was 13 ±
8.5% after 2 years and 18 ± 2.4% after 3 years comparing with the first year value.
Therefore, one possible explanation for the organic carbon decrease is the organic
degradation combined with the leaching of a small part of the BCHyPy fractions in the
biochar amended soil after four years of weathering.
5.1.3.3. Labile fraction
The non-stable fraction is also very important and, in addition to the quantification of
the BCHyPy fraction, HyPy also allows the molecular characterisation of the labile fraction
defined as non-BCHyPy fraction. The labile fraction is the part of the biochar that during its
storage in soil is released by predominantly microbial activity within the first few weeks or
months after the application of biochar (Masek et al., 2013). Therefore, the labile fraction
that evolves from biochar during its storage in soil is highly likely to impact on microbial
106
activity, and therefore affects the functioning of the soil as a whole. Some studies have
examined the non-BCHyPy fraction from soils (Meredith et al., 2013). These fractions are
likely to be dominated by products of decomposition of labile organic matter such as
lignocellulosic material and humic acids. However, there are not researches that identifie
labile organic compounds in biochar amended soils. Moreover, the HyPy allows to
characterize these materials with better preservation than typically encountered with more
traditional pyrolytic methods.
5.1.3.4 Aromatic hydrocarbon non-BCHyPy fraction
As well as isolating the BCHyPy fraction, HyPy also allows the characterisation of the
non-BCHyPy material at a molecular level by GC-MS analysis. The remaining material that
was labile under HyPy conditions, as so is defined as non-BCHyPy can be recovered and
characterised. The mass chromatograms of the non-BCHyPy fraction (Fig. 5.1.2) derived
from soil and biochar amended soil contain a high abundance of PAHs, predominantly
fluoranthene and pyrene. The PAHs released and trapped following HyPy treatment can be
considered as part of the BC continuum. Their presence in the non-BCHyPy fraction will be
due to their greater volatility relative to the larger more condensed and refractory aromatic
domains which form the BCHyPy.
107
Fig. 5.1.2. Total ion chromatograms from HyPy of biochar amended soil and soil (nonBCHyPy fraction).
Non-BCHyPy
Biochar amended soil
nC16
nC18
TTB
i.s.
Non-BCHyPy
Soil
nC16
nC18
TTB
Std
108
Table 5.1.3. Concentration of PAHs (µg g-1) released by the HyPy of biochar amended
soil.
Biochar amended soil
PAHs
Aug 2011
Dec 2011
May 2012
May 2013
Naphthalene
0.15±0.32
n.d.
n.d.
n.d.
Acenaphthene
0.33±0.32
n.d.
n.d.
n.d.
Fluorene
0.13±0.06
0.10±0.13
0.09±0.02
0.08±0.12
1.4±1.1
1.7±1.7
0.89±0.68
0.91±0.42
0.05±0.03
0.05±0.02
0.08±0.03
0.14±0.07
3.0±1.5
3.3±1.4
2.9±0.9
2.4±1.1
33.2±17.6
36.3±15.5
38.0±14.3
30.3±12.8
Chrysene
n.d.
n.d.
n.d.
n.d.
Benzo[a]anthracene
n.d.
n.d.
n.d.
n.d.
0.43±0.12
0.56±0.34
0.55±0.39
0.43±0.06
Benzo[b]fluoranthene
n.d.
n.d.
n.d.
n.d.
Benzo[k]fluoranthene
n.d.
n.d.
n.d.
n.d.
Benzo[a]pyrene
n.d.
n.d.
0.02±0.04
n.d.
Indeno[1,2,3-cd]pyrene
n.d.
n.d.
0.15±0.33
n.d.
Dibenzo[a,h]anthracene
n.d.
n.d.
n.d.
n.d.
Benzo[ghi]perylene
0.29±0.25
0.40±0.37
0.30±0.21
0.18±0.23
Total PAHs
38.9±20.5
42.4±19.3
42.9±16.4
34.5±14.3
Phenanthrene
Anthracene
Fluoranthene
Pyrene
Methylchrysene
Notes: Values in the tables are the mean value ± standard deviation from five replicates.
109
Table 5.1.4. Concentration of PAHs (µg g-1) released by the HyPy of soil.
Soil
PAHs
Aug 2011
Dec 2011
May 2012
May 2013
Naphthalene
n.d.
n.d.
n.d.
n.d.
Acenaphthene
n.d.
n.d.
n.d.
n.d.
Fluorene
0.13±0.02
0.13±0.05
n.d.
n.d.
Phenanthrene
0.75±0.17
0.44±0.13
0.27±0.07
0.22±0.10
Anthracene
0.09±0.05
0.09±0.08
0.05±0.01
0.06±0.02
Fluoranthene
1.84±0.53
1.57±0.62
1.55±0.18
1.51±0.32
Pyrene
20.5±6.3
17.5±6.6
15.9±2.2
16.7±4.2
Chrysene
n.d.
n.d.
n.d.
n.d.
Benzo[a]anthracene
n.d.
n.d.
n.d.
n.d.
0.27±0.11
0.39±0.21
0.30±0.04
0.13±0.08
Benzo[b]fluoranthene
n.d.
n.d.
n.d.
n.d.
Benzo[k]fluoranthene
n.d.
n.d.
n.d.
n.d.
Benzo[a]pyrene
n.d.
n.d.
n.d.
n.d.
Indeno[1,2,3-cd]pyrene
n.d.
n.d.
n.d.
n.d.
Dibenzo[a,h]anthracene
n.d.
n.d.
n.d.
n.d.
Benzo[ghi]perylene
0.17±0.15
0.18±0.08
0.11±0.09
n.d.
Total PAHs
23.9±7.2
20.6±7.8
18.2±2.4
18.7±4.4
Methylchrysene
Notes: Values in the tables are the mean value ± standard deviation from five replicates.
The PAHs detected and quantified in amended and untreated soils ranged from 2-ring
compounds (naphthalene) to 6-ring compounds (benzo[ghi]perylene), with the 4-ring
compound pyrene being the most abundant (Table 5.1.3 and Table 5.1.4). In almost all of
the soils, fluorene, phenanthrene, anthracene, fluoranthene and pyrene were detected. This
range of ring size is consistent with the PAHs distribution found in the non-BCHyPy fraction
generated by the HyPy of soil samples (Meredith et al., 2013), and the definition of BC HyPy
as being composed of PAHs with >7 rings proposed by Meredith et al. (2013). However, in
Meredith et al. (2013) has been reported a bigger average ring distribution.
110
We found that biochar amendment influence soil PAHs concentration during the
incubation period (Fig. 5.1.3). The total mean concentration value of PAHs in the amended
soils resulted higher than those of untreated soils during the 4 years of experiment (Fig.
5.1.3). The total PAHs concentration in untreated soils ranged between 10.4 and 31.9 µg g1
and in biochar amended soils between 16.1 and 75.2 µg g-1 (Table 5.1.3 and Table 5.1.4).
In addition, in the amended soils the concentrations of the PAHs did not decrease during
the four years following biochar application (Fig. 5.1.3). However, the differences were
not statistically significant due to the high dispersion of PAH values between samples
withdrawn from the same parcel (n = 5) (Table 5.1.3).
Figure 5.1.3. Concentration of PAHs (µg g-1) released by the HyPy of soil and biochar
amended soil
PAHs concentration (μg g-1)
70
60
50
40
Biochar amended soil
30
Soil
20
10
0
Aug 2011
Dec 2011
May 2012
May 2013
A detailed analysis of the contribution of the individual PAHs in biochar not subject to
environmental degradation and biochar amended soil indicated a similar distribution
profile in the non-BCHyPy fraction, with dominance of fluoranthene and pyrene in all the
samples studied (Table 5.1.3 and Table 5.1.5). However, also in the control soils,
fluoranthene and pyrene dominated the PAH profiles, supplying 7.6 ± 2% and 85 ± 8% of
the total PAH concentrations, respectively. Therefore, the distribution profiles of PAHs,
which comprise approximately 5-10% by weight of the non-BCHyPy, do not reflect biochar
modification.
Moreover, these long term field trials did not allowed to observe systematic changes in
the PAH distribution of the non-BCHyPy fractions. Four years after the addition of biochar
111
to agricultural soils, the PAHs not were degraded and the distribution profile is the same
during the 4 years of experiment.
Table 5.1.5. Observed concentration of PAHs released by the HyPy of biochar and a
corresponding biochar amended soil (2.22% (w/w) of reference biochar ).
Biochar
(µg g−1)
PAHs
Naphthalene
a
Soil + biochar
(µg g−1)
n.d.
n.d.
Acenaphthene
4.7±0.6
0.24
Fluorene
17±3.5
0.30
419±79.0
5.45
10±1.7
0.19
Fluoranthene
149±23.3
3.41
Pyrene
468±110
23.28
Chrysene
n.d.
n.d.
Benzo[a]anthracene
n.d.
n.d.
7.0±1.6
0.35
Benzo[b]fluoranthene
n.d.
n.d.
Benzo[k]fluoranthene
n.d.
n.d.
Benzo[a]pyrene
n.d.
n.d.
Indeno[1,2,3-cd]pyrene
n.d.
n.d.
Dibenzo[a,h]anthracene
n.d.
n.d.
Benzo[ghi]perylene
n.d.
0.13
1075±217
33.2
Phenanthrene
Anthracene
Methylchrysene
Total PAHs
Notes: Values in the tables are the mean value ±1 standard deviation from five replicates.
The last column reports the relative percent difference between the measured and expected
value. aThe expected value is the concentration calculated from the PAH concentration
obtained by summing the soil and biochar contributions.
112
5.1.3.Aliphatic hydrocarbons non-BCHyPy fraction
Fig. 5.1.2. shows mass chromatograms of the non-BCHyPy fraction derived from soil
and biochar amended soil, which was found to contain, in addition to PAHs, a high
abundance of n-alkanes, predominantly even numbered homologs. The n-alkanes comprise
approximately 10-20% by weight of the non-BCHyPy fraction.
The concentrations and diagnostic indices of n-alkanes in non-BCHyPy fraction of the
soil and biochar amended soil are presented in Table 5.1.6. and Table 5.1.7. The labile
fraction generated by the HyPy of soil (Fig. 5.1.4.) and biochar amended soil (Fig. 5.1.5) is
dominated by n-alkanes in the range nC13 to nC27 (the low carbon number compounds
having been lost to evaporation), with a distribution having maxima at nC16 to nC18, and a
even/odd predominance (C14 to C26 homologues CPI = 0.21 soil, CPI = 0.23 amended soil).
The total concentrations of n-alkanes in soil were higher than ones in the biochar amended
soil during the 4 years of experiment (Fig. 5.1.6). In untreated soils concentrations of nalkanes (nC13 to nC27) ranged from 64.2 µg g-1 to 130.7 µg g-1 and in amended soils from
18.2 µg g-1 to 72.9 µg g-1 (Table 5.1.6. and Table 5.1.7). After almost one year following
biochar application, the total mean concentration values of n-alkanes in biochar amended
soils was approximately 3 times higher than the control soil, both in August (37.2 vs. 106.7
µg g-1) and December (44.4 vs. 102.6 µg g-1) (Fig. 5.1.6).
Figure 5.1.4. Soil GC-MS mass chromatograms. Reconstructed ion chromatogram m/z 57
showing the n-alkane distribution present in the non-BCHyPy fraction.
nC17
nC16
Non-BCHyPy
Soil
nC18
m/z 57
nC19
nC20
nC15
nC21
nC22
nC14
nC22
nC24
nC26
nC13
113
i.s.
Fig. 5.1.5. Biochar amended soil GC-MS mass chromatograms Reconstructed ion
chromatogram m/z 128 + 154 + 166 + 178 + 202 + 231 + 242 + 276 showing the major
PAHs present in thenon-BCHyPy fraction.
m/z 128+154+166+178
+202+231+242+276
Non-BCHyPy
Biochar amended
soil
i.s.
Figure 5.1.6. Concentration of n-alkanes (µg g-1) released by the HyPy of soil and biochar
amended soil.
n-alkanes concentration μg g-1
140
n-alkanes
120
100
80
Biochar amended soil
60
Soil
40
20
0
Aug 2011
Dec 2011 May 2012 May2013
114
The characteristic relative distribution of n-alkanes in soil (Fig. 5.1.4) is not affected by
biochar modification. The effect of biochar on soil reduces the n-alkanes concentrations,
but both biochar amended soils and untreated soils are dominated by the same aliphatic
compounds. In particular, n-alkanes are characterized by a higher abundance of medium
chain homologues (nC16, nC18, nC20, nC22 and nC24) and are dominated by nC16 and nC18.
The Carbon Preference Indices (CPI) which expresses the ratio of odd-carbon
numbered to even-carbon-numbered n-alkanes is useful to determine the degree of
biogenic versus petrogenic input (Simoneit and Mazurek, 1982; Simoneit, 1989; Zheng et
al., 2000; Young, 2002). (Mazurek and Simoneit, 1984). n-alkanes originate from
epicuticular waxes of terrestrial plants and exhibit high values of CPI (CPI >1), whereas
CPI values for vehicular emissions and other anthropogenic activities are close to unit (CPI
~1).
The calculated CPI values for soil (0.21) and for biochar amended soil (0.23) are
similar (Table 5.1.8) and comparable to those observed for soil dichromate residue in
Meredith et al. (2013). This even numbered distribution, with a CPI of the C14 to C24
homologues of 0.21 and 0.23 indicates the importance of anthropogenic activities. These nalkanes were not predominantly original constituents of the soil organic matter. The nalkanes found in terrestrial soils are commonly dominated by odd-carbon-numbered nalkanes in the nC23–nC33 range derived from the epicuticular waxes of higher plants (Zelles
et al., 1999). However, a more probable source is from biolipids which exhibit an even/odd
preference, and typically include a high abundance of the C16 homologue, hexadecanoic
(palmitic) acid, in the short-chained (C10 to C20) fraction which is predominantly derived
from microbial biomass. The acidic species are known to be hydrogenated under HyPy
conditions to form the corresponding even-numbered n-alkanes (Meredith et al., 2006).
115
Table 5.1.6. Concentration of n-alkanes (µg g-1) released by the HyPy of biochar amended
soil.
Biochar amended soil
n-alkanes
Aug 2011
Dec 2011
May 2012
May 2013
nC13
0.52±0.26
0.003±0.002
0.003±0.001
0.024±0.003
nC14
2.8±1.0
0.037±0.017
0.002±0.001
1.04±10.02
nC15
3.5±0.9
1.62±0.9
0.19±0.11
2.8±1.7
nC16
14±5.5
11.3±6.9
7.6±5.3
16±6.7
nC17
2.8±1.2
2.9±0.9
3.4±1.6
3.2±1.1
nC18
12±6.9
18±6.9
20±10
14±5.8
nC19
0.78±0.38
1.4±0.5
1.9±0.9
1.1±0.4
nC20
1.73±1.18
3.2±1.3
4.1±2.6
2.5±1.1
nC21
0.46±0.29
0.81±0.30
1.08±0.7
0.68±0.31
nC22
1.0±1.0
2.9±1.3
3.98±1.9
2.46±1.24
nC23
0.28±0.21
0.60±0.37
0.71±0.55
0.51±0.28
nC24
0.61±0.50
1.1±0.8
1.34±1.42
1.02±0.48
nC25
0.11±0.07
0.14±0.06
0.24±0.20
0.21±0.05
nC26
0.13±0.10
0.16±0.09
0.30±0.26
0.27±0.10
nC27
0.05±0.03
0.17±0.07
0.16±0.15
0.17±0.13
Total nC13-nC27
40.5±17.8
44.4±11.6
45.1±24.0
46.1±18.6
CPI nC12-nC24 a
0.26
0.20
0.23
0.20
a
Carbon preference index (CPI) formula of Bray and Evans (1961).
116
Table 5.1.7. Concentration of n-alkanes (µg g-1) released by the HyPy of soil.
Soil
n-alkanes
Aug 2011
Dec 2011
May 2012
May 2013
nC13
0.59±0.07
0.91±0.58
0.004±0.003
0.064±0.06
nC14
2.6±0.95
3.1±2.1
0.025±0.014
1.1±0.7
nC15
5.3±1.3
5.7±1.2
1.1±0.3
3.0±0.9
nC16
30±5.6
26±2.0
18.8±0.4
25±5.1
nC17
5.3±3.0
5.6±0.8
6.06±0.5
6.1±1.5
nC18
34±8.9
29±3.6
36±2.4
35±11.4
nC19
2.5±0.4
2.1±0.7
2.99±0.3
2.42±1.1
nC20
4.2±3.6
8.0±1.4
9.78±0.8
7.2±3.3
nC21
2.38±0.6
2.4±0.5
3.04±0.4
1.7±0.9
nC22
10±4.2
11±3.1
14±2.3
7.6±4.4
nC23
2.2±1.0
2.1±0.8
2.2±1.3
1.4±0.9
nC24
4.8±2.7
4.9±2.5
6.3±1.1
2.8±1.8
nC25
0.73±0.41
0.66±0.42
0.92±0.19
0.41±0.29
nC26
1.11±0.72
1.01±0.65
1.27±0.3
0.56±0.38
nC27
0.37±0.23
0.32±0.20
0.47±0.33
0.17±0.10
Total nC13-nC27
107±16.7
103±17.7
102±8.0
93.8±18.6
CPI nC12-nC24 a
0.22
0.24
0.19
0.19
a
Carbon preference index (CPI) formula of Bray and Evans (1961).
5.1.4. Conclusions
A short-term incubation study was carried out to investigate the effect of biochar
addition to soil on the level of organic carbon and to study the persistence and resistance of
biochar in the environment. Although the carbon added with biochar should be quite
recalcitrant to degradation, it is well known that a variable component of biochar is labile
(degradable on annual/decadal timescales) and hence, only a proportion of total carbon in
biochar provides long-term carbon sequestration.
117
Previous studies reported that HyPy is potentially a precise method for BC
measurements in soil, lignocellulosic material, coals, and petroleum source rocks. The
results presented in this study suggest that HyPy provides a rapid and convenient technique
for the quantification of BC and determination of the stable carbon in soil treated with
biochar. The ability of the method to determine BC in the biochar amended soil was
evaluated. The findings of this study showed that biochar amendment significantly
influence soil BC concentration during the incubation period. In particular, the obtained
concentrations of BC in the amended soil are significantly higher than that in the untreated
soil. Obviously, the effect of biochar addition in soils on the level of BC will depend on the
BC in the original biochar. The high BCHyPy value found for biochar (83±3.3%) used in this
study suggests that it should be quite recalcitrant to degradation.
Moreover, the HyPy allowed the characterisation on a molecular level of labile fraction
defined as non-BCHyPy fraction. In addition to a number of PAHs, the non-BCHyPy fraction
was also found to contain a significant abundance of n-alkanes, with a marked
predominance of even-numbered homologues. These compounds are probably derived
from lipids, hydrogenated during HyPy. However, further researches are need to
characterise the BCHyPy residues and more fully confirm that they are entirely free of nonBC material.
118
5.2. Characterization of biochar stability by hydropyrolysis and
pyrolysis-GC/MS
5.2. Biochar stability characterization by hydropyrolysis and pyrolysis-GC/MS
5.2.1. Introduction
Biochar is carbonaceous solid formed by the pyrolysis of biomass which attracts
research interest due to its potential value for long-term carbon sequestration. The addition
of biochar to soil has been proposed as strategy that not only sequesters carbon in soils but
also at the same time mitigates different environmental issues. Research has demonstrated
that biochar has considerable potential as a sustainable tool for carbon sequestration, soil
amelioration, greenhouse gas emissions reduction and fertilizer runoff reduction, as well as
waste management (Glaser et al., 2002; Lehmann et al., 2009; Sohi et al., 2010; Woolf et
al., 2010; Zavalloni et al., 2011; Galinato et al., 2011; Kookana et al 2011; Yao et al.,
2012).
A key requirement for the use of biochar as tool for environmental management is that
the carbon in the biochar is stable, meaning that a substantial fraction of the carbon
sequestered is not re-mineralized on at least centennial timescales (Gurwick et al., 2013).
However, a variable component of the carbon in many biochars is degradable on annual to
decadal timescales and hence, only a proportion of total carbon in biochar provides longterm carbon sequestration (Bird et al., 1999; Zimmermann et al., 2012). Although our
understanding of biochar carbon stability has improved in recent years (Ameloot et al.,
2013), there is limited research on process conditions to produce a biochar suitable and
highly stable for the long-term carbon sequestration (Conti et al., 2014; McBeath et al.,
2015). The properties of biochar, including stability, depend on the type of feedstock,
pyrolysis temperature and pyrolysis method (Labbe et al., 2006; Nguyen and Lehmann,
2009; Singh et al., 2012). However, biochar stability depends also on the environmental
factors (temperature, rainfall, soil type) of the site where the biochar is incorporated into
the soil (Czimczik and Masiello, 2007).
A number of approaches have been proposed to assess biochar stability. However, there
is no agreed methodology for determining the long-term stability of biochar yet. The
methods which have been proposed to assess biochar stability include solid state nuclear
119
magnetic resonance spectroscopy (solid state
13
C NMR) (McBeath and Smernik, 2009),
Fourier transform infrared spectroscopy (FTIR) (Michel et al., 2009), proximate analysis
using the fixed carbon as a measure of stability (ASTM Standard D3175; 2007), thermal
analysis (thermogravimetry, TG; de la Rosa et al., 2008), molecular markers by means of
pyrolysis-gas chromatography-mass spectrometry (Py-GC-MS; Kaal et al., 2008, 2009,
Conti et al., 2014), benzene polycarboxylic acid method (Brodowski et al., 2005), O:C or
H:C molar ratios (Spokas, 2010; Enders et al., 2012; IBI Guidelines, 2012) and chemical
oxidation (Cross and Sohi, 2013). Many studies reported that chemical oxidants, such as
K2Cr2O7, KMnO4, HNO3, and H2O2, could be used to evaluate the oxidative nature of
biochar and reflect the long-term stability of biochar (Trompowsky et al., 2005; Knicker et
al., 2007; Calvelo Pereira et al., 2011; Li et al., 2014). In Masek et al. (2013) stable carbon
in biochar was determined using an accelerated ageing assay. This assay involved the
thermal and chemical oxidation of milled biochar samples. Samples were placed in 5%
hydrogen peroxide and heated to 80 °C, and carbon stability then was calculated
gravimetrically using the %C data of samples before and after oxidation. This approach is
considered to be more representative of the degradation processes to which biochar would
be subjected in the environment. The results of the accelerated ageing experiments have
demonstrated for a range of biochars chemical behaviours consistent with the hypothesis
that biochars produced at higher temperatures exhibit more resistance to oxidative
degradation carbon (black carbon) fractions in biochar. Black carbon (BC) is an important
component of organic carbon (OC), and it is defined as the carbon-rich (>60%) product of
the incomplete combustion of fossil fuels and biomass (Goldberg, 1985), that includes a
range of products such as char, charcoal, ash, and soot (Preston and Schmidt, 2006).
An emerging pyrolytic approach isolating and quantifying BC in a range of
environmental matrices is hydropyrolysis (HyPy) combined with GC-MS (Ascough et al.,
2009; Meredith et al., 2012; Wurster et al., 2013). These studies have shown that the HyPy
method removes all labile organic matter (defined as non-BCHyPy) (Wuster et al., 2012), so
isolating a highly stable portion of the BC (BCHyPy) predominantly composed of >7 ring
aromatic domains (Meredith et al., 2012). The high hydrogen pressure and slow heating
rate employed, together with the presence of a sulphided molybdenum catalyst, prevent the
generation of secondary char (Love et al., 1995) encountered with other chemical or
thermal oxidative methods. In addition to the quantification of the BCHyPy fraction, HyPy
120
also allows the molecular characterisation of the biochar labile fraction defined as nonBCHyPy fraction.
The molecular signature of the thermally labile fraction of biochar was also examined
by pyrolysis-GC/MS and its association to the stability of the carbon in biochar
investigated (Pereira et al., 2011; Conti et al., 2014). In particular, the GC-MS traces
(pyrograms) of biochar were featured by peaks associated with benzene, toluene,
naphthalene, biphenyl, dibenzofuran and benzonitrile (Kaal et al., 2009). These pyrolysis
products were assumed to represent the charred fraction abundant of aromatic structures
that occur in a thermally labile form in the carbonaceous matrix. In addition, pyrolysis
product ratios representing the relative abundance of alkylated and parent compounds (e.g.,
benzene/toluene peak area ratio) were proposed as indicators for the presence of saturated
alkyl bridges between polyaromatic structures and hence a measure of the charring
intensity (Kaal et al., 2012). Therefore, Py-GC-MS is able to provide molecular indices of
biochar stability. However, there are no studies aimed at comparing these indices to those
arising from HyPy.
The aim of the present study is to assess the impact of production conditions on biochar
stability, providing moreover a comparison between molecular analysis by Py-GC-MS and
HyPy on biochar samples produced from three feedstock and the same pyrolysis unit.
Different process conditions, charring temperature and residence time, were utilised to
obtain biochars with different degrees of charring.
5.2.2. Experimental section
5.2.2.1. Samples
Three different types of biomass were used as feedstock materials: pine wood chips –
with an average size of 3 cm × 2 cm × 0.5 cm – from Robeta Holz OHG, Milmersdorf,
Germany; beech wood spheres – with a diameter of 25 mm –, provided by Meyer and
Weigand Gmbh, Nordlingen, Germany; corn digestate derived from maize silage.
Biochar samples were produced by pyrolysis of sample using a stainless steel fixed-bed
reactor of 102.5 cm height and 22 cm of internal diameter. The inert atmosphere is
provided by a N2 flow entering the reactor (20 L min-1 and 50 L min-1) from the bottom
through a stainless steel grate to get an uniformly distributed flow. The samples (in the
range of kilograms) were uniformly placed inside the reactor in a stainless steel container
121
of 21 cm of diameter and 56 cm height which is placed directly on the previously
mentioned grate. The reactor is externally heated with a wire heater with a maximum
power of 3000 W placed on the external reactor wall. Both flanges in the reactor are also
heated and insulated to reduce heat losses. The N2 flow is preheated before entering the
reactor as well. The temperature operation of this preheater is 600 °C.
Pyrolyses were performed at three different temperatures, 340 °C, 400 °C and 600 °C.
The biochar samples obtained were labeled as PW ID 1, PW ID 2, BW ID 1, BW ID 2, CD
EU 1 and CD EU 2 (where PW, BW, CD stand for pine wood, beech wood and corn
digestate, respectively; 1 and 2 indicated the highest and lowest pyrolysis temperatures,
respectively; ID and EU, see Table 5.2.1).
Table 5.2.1. Biomass feedstock and pyrolysis conditions of biochar samples. (# : Sample
identifiers).
PW ID 1
Raw
material
Pine wood
Volatiles
%
10.30
Max T
°C
600
N2 flow
L/min
20
PW ID 2
Pine wood
33.70
400
20
CD EU 1
Corn digestate
12.66
600
20
CD EU 2
Corn digestate
15.93
400
20
BW ID 1
Beech wood
9.93
600
20
BW ID 2
Beech wood
-
340
50
#
5.2.2.2. Biochar bulk characterization
The pH of the biochar samples was determined by adding biochar to deionized water at
1:10 wt/wt mass ratio and pH measured at room temperature with a digital pH meter (HI
98103, Checker®, Hanna Instruments). Elemental composition (HCNS) was determined
by combustion using a Thermo Scientific Flash 2000 series analyzer. Ash was determined
as the residual mass left after exposure at 600 °C for 5 hours. The oxygen content was
calculated from the mass balance: Oxygen (%) = 100 - Ash content (%) - C (%) – H (%) N (%). Moisture contents were determined (ASTM D-3173) at 105 °C.
122
5.2.2.3. Polycyclic aromatic hydrocarbons (PAHs)
The content of PAHs in biochars was measured in triplicate as described in Fabbri et
al., (2013), but using 16 PAHs surrogate of each of the 16 US EPA PAHs instead of 3
PAHs
surrogate.
acenaphthene,
The
measured
fluorene,
benzo(a)anthracene,
PAHs
phenanthrene,
chrysene,
included
naphthalene,
anthracene,
benzo(b)fluoranthene,
acenaphthylene,
fluoranthene,
pyrene,
benzo(k)fluoranthene,
benzo(a)pyrene, dibenz(ah)anthracene, benzo(ghi)perylene, and indeno(1,2,3-c,d)pyrene.
Briefly, about 0.5 g of biochar was spiked with 0.1 mL of a 5 mg l-1 solution of
surrogate 16 EPA PAHs (prepared from Dr. Ehrenstorfer PAH-Mix 9 deuterated, 10 ng µl1
) and soxhlet extracted with acetone/cyclohexane (1:1, v/v) for 36 hours. The solution was
filtered, added with 1 ml of n-nonane (keeper), carefully evaporated by rotatory vacuum
evaporation at 40 °C and cleaned up by solid phase extraction onto a silica gel cartridge
before analysis with a Agilent HP 6850 GC coupled to a Agilent HP 5975 quadrupole mass
spectrometer; GC-MS conditions were those detailed in Fabbri et al., (2013). Recovery of
surrogate PAHs was determined with respect to the internal standard tri-tert-butylbenzene.
Results are reported as averages of three replicates analyses.
5.2.2.4. Pyrolysis-GC/MS
Py-GC-MS analyses were performed using an electrically heated platinum filament
CDS 1000 pyroprobe valved interfaced to a Varian 3400 GC equipped with a GC column
(HP-5-MS; Agilent Technologies 30 m × 0.25 mm, 0.25 μm) and a mass spectrometer
(Saturn 2000 ion trap, Varian Instruments) set at an electron ionization at 70 eV in full
scan acquisition (10–450 m/z). A quartz sample tube containing of weighed biochar sample
(5-10 mg) added with 1 µL of internal standard solution (o-isoeugenol at 1000 mg L-1 in
methanol) was inserted into the Py-GC interface (300 °C) and then pyrolysed at 900 °C
(set temperature) for 100 s with helium as carrier gas (100 ml min-1). The following
thermal program was used: 35 °C to 310 °C at 5°C min-1.
Yields were estimated from the ratio of the peak area integrated in the mass
chromatogram of a characteristic ion of the selected pyrolysis product and the peak area of
the internal standard, the quantity of added internal standard and the amount of sample
pyrolysed (Torri et al., 2010). An unitary relative response factor was assumed for all the
quantified compounds on the basis that our objective was the comparison between samples
123
on a quantitative base rather than the knowledge of the absolute yield of each pyrolysis
product. Total yields were the summed yields of all the selected pyrolysis products.
A set of 38 pyrolysis products among the most abundant and representative of
biological precursors was selected on the basis of a previous work (Fabbri et al., 2012)
(Table 5.2.2). On each biochar sample was carried out a single analysis and the pyrolysis
products were quantified both in terms of yields (µg g-1) and relative abundance
considering a relative standard deviation (RSD) between 8-36 % that is typical of Py-GCMS analysis.
Table 2. Pyrolysis products of biochar, the mass to charge ratio (m/z) of the quantitation
ion and their predominant origin: C, charred biomass; H holocellulose (sugars); L, lignin;
P, proteins (nitrogen-containing compounds).
# Pyrolysis product
m/z origin # Pyrolysis product
m/z origin
1
2
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
78
75
96
67
91
98
91
98
91
104
91
94
118
103
117
108
107
109
132
122
122
122
122
128
110
142
142
138
150
154
154
137
168
164
178
178
202
202
Benzene
Hydroxyacetone
Dimethylfuran
Pyrrole
Toluene
2-Methyltiophene
o-Xylene
Fufurilic alcool
m/p-Xylene
Styrene
Ethylbenzene
Phenol
Benzofuran
Benzonitrile
Indole
3-methylphenol
4-methylphenol
Guaiacol
Methyl-benzofurans (3 isomers)
C
H
H
P
C
P
C
H
C
C
C
L
C
C
P
L
L
L
C
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
2-ethylphenol
2,5-dimethylphenol
2,3-dimethylphenol
3-ethylphenol
Naphthalene
Catechol
2-methylnaphthalene
1-methylnaphthalene
4-methylguaiacol
4-vinylguaiacol
Syringol
Biphenyl
4-ethylguaiacol
4-methylsyringol
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
L
L
L
L
C
L
C
C
L
L
L
C
L
L
C
C
C
C
C
5.2.2.5. Hydropyrolisis
Hydropyrolysis (HyPy) tests were performed using the procedure described in detail in
a number of publications (e.g. by Ascough et al., 2009; Meredith et al., 2012). Briefly, 50100 mg of biochar sample were loaded with a Mo catalyst using an aqueous/methanol 0.2
124
M solution of ammonium dioxydithiomolybdate [(NH4)2MoO2S2]. Catalyst weight was ~
10% of the sample weight. The catalyst loaded biochar samples were placed within
shortened borosilicate pipette ends (20 mm long), plugged at each end with pre-cleaned
quartz wool and then placed in the HyPy reactor. We used the recommended temperature
program previously optimized for pyrogenic carbon quantification where the samples are
heated at rate of 300°C min-1 from 50 to 250°C, then heated at 8 °C min -1 from 250°C until
the final temperature of 550°C for 2 min (Ascough et al., 2009; Meredith et al., 2012), all
under a hydrogen pressure of 15 MPa. A hydrogen sweep gas flow of 5 L min-1, measured
at ambient temperature and pressure, ensured that the products were quickly removed from
the reactor vessel, and subsequently trapped in a silica gel-filled trap cooled by dry ice.
5.2.2.6. Black carbon quantification
The BC (reported as BCHyPy) content of each sample was derived by comparing the
organic carbon (OC) content of the catalyst loaded samples prior to HyPy with those of
their HyPy residues (Eq. (1)). Elemental composition (HCNS) was determined by
combustion using a Thermo Scientific Flash 2000 series analyzer.
BCHyPy (BC=OC%) =
Residual OC (mg C in HyPy residue including spent catalyst)
Initial OC (mg C in sample including catalyst)
x 100
(1)
5.2.2.7. Non-BCHyPy fraction characterisation
The non-BCHyPy fraction (hydropyrolysate) from the soil, biochar amended soil and
biochar samples were desorbed from the silica recovered from the trap with 10 ml aliquots
of n-hexane and dichloromethane (DCM). The eluents were evaporated to 1 ml at room
temperature for 12 h prior to analysis. GC–MS analyses in full scan mode (m/z 35-650)
were performed on 6850 Agilent HP gas chromatograph connected to a 5975 Agilent HP
quadrupole mass spectrometer (EI mode, 70 eV), equipped with an autosampler and a
split/splitless injector. Analytes were separated by a HP-5MS fused silica capillary column
(stationary phase poly[5% diphenyl/95%dimethyl]siloxane, 30 m × 0.25 mm i.d., 0.25 mm
film thickness), using helium as the carrier gas, and an oven programme of 50°C (hold for
2 min) to 300°C (hold for 33 min) at 5°C min-1. Samples (1 µl) were injected under
splitless conditions (1 min, then split ratio 1:50 to the end of analysis) with an injector
125
temperature of 280°C. For the quantification of PAHs the mass chromatograms of the
molecular ion of each compound was used, following the addition of 100 µl of 1,3,5-tritert-butylbenzene (TTB, 100 mg l−1 Sigma-Aldrich) as internal standards, assuming a
response factor for each compound of 1.
The PAHs were identified by matching the retention times of each peak in the sample
chromatogram with those of a standard solution. Interfering coelution problems were
evaluated in the samples by comparing mass spectra of the samples with those of the
standards as well as with those from the NIST mass spectra library (NIST MS Search r.
2.0).
5.2.2.8. Statistical Analysis
Quantitative data are presented as mean values ± standard deviation (n = 2). Recovery
of surrogate PAHs was (mean ± %RSD for all the data set): 80% ± 6% naphthalene-d8,
69% ± 27% acenaphthylene-d8, 91% ± 4% acenaphthene-d10, 88% ± 21% fluorene-d10,
90% ± 17% phenanthrene-d10, 70% ± 23% anthracene-d10, 88% ± 10% fluoranthene-d10,
87% ± 9% pyrene-d10, 83% ± 18% chrysene-d12, 84% ± 10% benzo(a)anthracene-d12, 87%
± 11% benzo(b)fluoranthene-d12, 79% ± 15% benzo(k)fluoranthene-d12, 79% ± 23%
benzo(a)pyrene-d12,
75%
±
22%
indeno(1,2,3-c,d)pyrene-d12,
82%
±
12%
dibenz(ah)anthracene-d14 and 77% ± 19% benzo(ghi)perylene-d12. Student t tests were
conducted with Excel (2011) to evaluate significant difference between two parameters of
biochar. Linear (Pearson) correlation coefficient between two variables r(df), where df
stands for degrees of freedom, was determined for all the investigated parameters. Two set
of data were assumed to be correlated when the absolute value of r was larger than the
critical value at the level of significance p = 0.01 for two-tailed test.
5.2.3. Results and discussion
5.2.3.1 Biochar characteristic
Results of biochar characterizations are reported in Tables 5.2.3 and 5.2.4. The biochars
had a range of 46.4–91.6% carbon; 1.1–47.3% ash; 0.2–1.9% nitrogen; 3.8–22.9% oxygen.
The biochars showed profound differences in properties, depending on feedstock and
pyrolysis temperature (Table 5.2.3). Pyrolysis temperature showed significant effect on
126
elemental compositions of wood biochars and to a lesser extent on that of corn digestate
biochars. In particular, the data showed that carbon content of wood biochar increased with
temperature, while the oxygen and hydrogen contents decreased. This resulted in lower
H/C and O/C atomic ratio values at increasing final temperature (Table 5.2.3).
The degree of carbonisation of chars is generally expressed by molar H/C (Calvelo Pereira
et al., 2011; Enders et al., 2012) or O/C ratios (Spokas et al., 2010; Brodowski et al., 2005).
The O/C ratios here investigated chars ranged from 0.04 (PW ID 1) to 0.24 (PW ID 2) in
accordance to the loss of oxygenated functionalities with increasing carbonisation (Krull et
al., 2009), and were strongly correlated with molar H/C ratios (R = +0.96, Table 5.2.3).
However the carbon and oxygen contents of beech and pine wood biochar was higher
compared to that of corn digestate biochar. On the contrary, the corn digestate biochar
showed higher ash and nitrogen contents. The differences in ash and carbon contents can
be linked to the chemical composition differences between wood and corn digestate. Wood
contains more cellulose and hemicelluloses and during high temperature pyrolysis (> 500
°C), the components are reduced to carbon thus the higher carbon content in wood biochar
(Ahamedna et al., 2000, Keiluweit et al., 2010, Al-Wabel et al., 2013).
The ash content of biochar samples was influenced mainly by feedstock and to a lesser
extent by pyrolysis temperature with ash content increasing with pyrolysis temperature.
The increase in ash content should result from progressive concentration of minerals and
destructive volatilization of lignocelluloses matters as temperature increased (Tsaia et al.,
2012). However, the ash content of corn digestate biochar was much higher (up to 47.27%)
than that in beech and pine wood (up to 1.51%).
127
Table 5.2.3. Elemental composition of biochar samples (# : Sample identifiers). Mean
values and % relative standard deviation (rsd) from two replicates.
#
H/C
(molar)
O/C
(molar)
C
(%)
H
(%)
N
(%)
O
(%)
Ash
(%)
Moisture
(%)
PW ID 1
0.33
0.04
91.6 ± 2.3
2.50 ± 0.069
0.21 ± 0.017
4.4 ± 2.4
1.32 ± 0.04
0.10 ± 0.001
PW ID 2
0.70
0.24
71.7 ± 0.81
4.20 ± 0.064
0.18 ± 0.011
22.9 ± 0.89
1.06 ± 0.02
0.19 ± 0.026
CD EU 1
0.22
0.06
46.4 ± 0.37
0.86 ± 0.019
1.58 ± 0.004
3.87 ± 0.43
47.27 ± 0.04
0.13 ± 0.014
CD EU 2
0.30
0.06
48.0 ± 0.82
1.21 ± 0.074
1.91 ± 0.048
3.80 ± 1.0
45.07 ± 0.05
0.08 ± 0.004
BW ID 1
0.29
0.08
87.6 ± 2.8
2.14 ± 0.18
0.19 ± 0.047
9.3 ± 2.0
1.51 ± 0.04
0.13 ± 0.006
BW ID 2
0.72
0.23
71.9 ± 2.4
4.33 ± 0.092
0.18 ± 0.005
22.4 ± 2.4
1.14 ± 0.08
0.18 ± 0.014
The concentrations of PAHs ranged between 2.2 (BW ID 2) and 18.9 µg g-1 (PW ID 2)
(Table 5.2.4. and Figure 5.2.1.), thus not all the biochars were below the levels
recommended by IBI or EBC (4-12 µg g-1). However, it is interesting to note that despite
the difference in feedstock at 600 °C the PAH levels were quite similar (2.2-2.9 µg g-1),
while were significantly different at low temperature (340-400°C). Therefore, the influence
of feedstock type on PAHs concentration is evidenced by results obtained from biochar
produced at low temperature. PAH levels tended to increase with decreasing H/C ratios,
indicating that a multitude of factors could influence the occurrence of PAHs in biochar
(Schimmelpfennig and Glaser, 2012). In fact, different trends were reported in the
literature (Hale et al., 2012; Keiluweit et al., 2012; Schimmelpfennig and Glaser, 2012),
such as decreasing or increasing PAH concentrations with increasing pyrolysis
time/temperature for slow and fast pyrolysis, respectively (Hale et al., 2012), or PAH
concentrations peaking at 500 °C in grass biochars produced in the 100-700 °C pyrolysis
interval (Keiluweit et al., 2012). For beech wood and corn digestate biochar, naphthalene
was the most abundant PAH, in accordance to previous studies (Hale et al., 2012; Hilber et
al., 2012; Schimmelpfennig and Glaser, 2012; Freddo, et al., 2012; Fabbri et al., 2013),
followed by phenanthrene. While for the pine wood biochar the most abundant PAH was
phenanthrene, followed by naphthalene in BW ID 1 and by fluoranthene and pyrene in BC
ID 2.
128
Table 5.2.4. Concentrations of the 16 US-EPA PAHs in biochar. Mean values in ng g-1
d.w. and standard deviation from two replicates.
Sample Id.
PAHs
Naphthalene
PW ID 1
PW ID 2
CD EU 1
-1
CD EU 2
-1
617 ± 8.1
423 ± 64
ng g
1512 ± 15
6039 ± 635
Acenaphthylene
49 ± 11
145 ± 29
243 ± 4.5
613 ± 52
326 ± 41
173 ± 31
Acenaphthene
119 ± 16
546 ± 24
144 ± 7.2
113 ± 16
176 ± 18
273 ± 57
Fluorene
273 ± 4.9
745 ± 2.4
124 ± 0.6
148 ± 6.5
292 ± 19
147 ± 22
Phenanthrene
778 ± 188
10244 ± 233
135 ± 19
130 ± 0.6
274 ± 37
183 ± 34
Anthracene
81 ± 7.1
824 ± 77
23 ± 23
22 ± 0.1
20 ± 2.3
23 ± 0.8
Fluoranthene
207 ± 46
3355 ± 135
61 ± 13
41 ± 0.3
74 ± 0.8
41 ± 4.2
Pyrene
45 ± 9.1
1732 ± 209
48 ± 9.3
44 ± 12
81 ± 4.3
48 ± 9.3
Chrysene
229 ± 12
181 ± 18
41 ± 6.6
12 ± 3.3
94 ± 7.8
6.4 ± 0.7
Benzo[a]anthracene
183 ± 24
341 ± 62
27 ± 5.9
23 ± 1.6
8.0 ± 0.5
20 ± 0.7
Benzo[b]fluoranthene
288 ± 30
197 ± 4.0
89 ± 18
33 ± 2.7
79 ± 15
32 ± 8.0
Benzo[k]fluoranthene
76 ± 15
37 ± 2.4
34 ± 3.0
35 ± 3.0
40 ± 6.8
12 ± 2.1
Benzo[a]pyrene
19 ± 2.6
25 ± 5.0
44 ± 7.7
20 ± 1.7
117 ± 11
79 ± 15
Indeno[1,2,3-cd]pyrene
n.d.
38 ± 6.0
n.d.
n.d.
n.d.
n.d.
Dibenzo[a,h]anthracene
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
Benzo[ghi]perylene
n.d.
70 ± 5.2
n.d.
n.d.
n.d.
n.d.
18905 ± 290
2524 ± 41
7274 ± 710
2730 ± 14
2240 ± 45
Σ 16 EPA PAHs
ng g
BW ID 1
BW ID 2
-1
ng g
2964 ± 290
129
1132 ± 12
1204 ± 3.2
Figure 5.2.1. Concentrations of the 16 US-EPA PAHs in biochar. Mean values in µg g-1
d.w. and standard deviation from two replicates.
25
PAHs solvent
µg g-1
20
15
10
5
0
PWID1 PWID2
BWID1 BWID2
CDEU1 CDEU2
5.2.3.2. Biochar carbon thermal stability stability determination via HyPy
It is important to determine how much of the carbon contained in biochar is potentially
stable over long periods of time as there are likely to be various fractions, differing in their
stability, ranging from very unstable (labile) fractions to very recalcitrant (stable) fractions.
The HyPy method has been demonstrated to remove almost all labile organic carbon,
leaving a highly stable residue with average polycondensed structures greater than 7 fused
rings (Meredith et al., 2012; Wuster et al., 2012). The low molecular weight non-BCHyPy
that are removed by HyPy along with any other residual labile organic compounds are
unlikely to be stable on centennial timescales due to their susceptibility to biological and
chemical oxidation (Knicker et al., 2008; Ascough et al., 2008).
Table 5.2.5 and Figure 5.2.2 show the results obtained from analysis of the biochar
samples using HyPy. It can be seen a significant difference in BCHyPy concentration and
mass losses during HyPy of the biochar samples. The feedstock source and pyrolysis
temperature clearly influenced the proportion of BC and the degree of condensation of
aromatic C (Table 5.2.5 and Figure 5.2.2). The wood biochars produced at 340-400 °C
contained the lowest proportions of BCHyPy fraction (15.7 – 20.5%), with the pine wood
biochar having lower proportions (15.7 ± 0.54%) than the biochar produced from beech
130
wood (20.5 ± 5.0%) (Table 5.2.5). The BCHyPy fraction of the 600 °C wood biochars
(Figure 5.2.2) contained the highest proportions of BCHyPy fraction (89.7-95.9%), with the
pine wood biochar again having higher proportions (95.9 ± 0.69%) than the biochar
produced from beech wood (89.7 ± 1.7%) (Table 5.2.5). Therefore, the fraction of BC as
proportion of total organic carbon, as determined by the HyPy method, increases from
15.7% at 400 °C to 95.9% at 600 °C for wood biochars, demonstrating that pyrolysis
temperature exerts a strong control on the formation of BC (Figure 5.2.2 and Table 5.2.5).
Table 5.2.5. Weight loss (%) during HyPy treatment and black carbon (BC) from HyPy of
the biochars. Mean values and standard deviation from two replicates.
Sample
PWID1
PWID2
BWID1
BWID2
CDEU1
CDEU2
BCHyPy (BC/OC)
%
sd
95.9
0.69
15.7
0.54
89.7
1.7
20.5
5.0
96.7
2.1
91.0
1.7
Weight loss
%
20.0
82.9
16.1
76.4
12.7
16.1
sd
2.2
0.11
3.2
3.1
3.2
4.0
The relatively low BCHyPy contents of the BW ID 2 and PW ID 2 biochars are to be
expected given that the temperature of formation for these samples was only 340 and 400
°C, respectively, which may have allowed for products of the incomplete thermal
degradation of cellulose and lignin to be preserved in biochar (Hammes et al., 2007). These
findings are consistent with those of some researches (McBeath and Smernik, 2009;
Nguyen et al., 2010; Masek et al., 2013) who also observed an increase in aromaticity and
aromatic condensation of biochars with increasing pyrolysis temperature. However, PW ID
2 and BW ID 2 biochars do not meet the quality criteria of the European Biochar
Certificate, which sets a range of minimum allowed threshold values for the black carbon
content in the biochars (10 - 40% of overall carbon).
The influence of feedstock type on the degree of carbonisation is evidenced by results
HyPy obtained from biochar of corn digestate. This feedstock at 400 °C produced a
thoroughly carbonised biochar, compared with BW ID 2 and PW ID 2. This was reflected
in (i) a low mass losses during HyPy; (ii) a high value of black carbon by HyPy; (iii) a low
value of non-BCHyPy fraction produced (Figure 5.2.2).
131
The BCHyPy content was inversely correlated with H/C and O/C ratios (R = -0.99 in
both cases, Table 5.2.3) confirming as biochars with low H/C and O/C values are graphitelike materials (i.e. soot, black carbon, activated carbon), they are expected to be more
stable and less prone to degradation (Masiello, 2004).
BC as proportion of OC (%)
Figure 5.2.2. Black carbon (BC) as proportion of organic carbon (OC) as measured by
HyPy of the biochar.
100
90
80
70
60
50
40
30
20
10
0
PW ID 1 PW ID 2 CD EU 1 CD EU 2 BW ID 1 BW ID 2
5.2.3.3. Biochar carbon thermal stability stability determination Py-GC-MS
The analysed biochar samples produced different pyrolysate patterns when subjected to
Py-GC-MS. Some typical pyrograms resulting from Py-GC-MS of biochar samples are
depicted in Fig. 5.2.3, the numbers in which refer to the pyrolysis product list (Table
5.2.2). Samples with low BCHyPy values produced complex pyrolysates with intense peaks
assignable to the pyrolysis products of hemicelluose, cellulose or lignin, on the contrary
samples with high BCHyPy ratios produced simple pyrograms with weak peaks of aromatic
hydrocarbons.
132
Figure 5.2.3. Total ion chromatograms from Py–GC–MS of biochar BW ID 2, BW ID 1,
PW ID 1, PW ID 2, CD EU 2 and CD EU 1. Peak attribution: (1), benzene; (2), toluene;
(3), m,p-xylene; (4), styrene; (5) ethylbenzene; (6), phenol; (7), indole; (8), 3methylphenol; (9). 4-methylphenol (10), guaiacol; (11), 2,5-dimethylphenol; (12), 2,3dimethylphenol; (13), naphthalene; (14), catechol. Internal standard: (i.s.) o-isoeugenol.
MCounts
1
2
8
2.0
1.5
7
3
1.0
i.s.
BW ID 2
4 5
0.5
MCounts
11 14
13
910 12
6
0.0
2.0
5
10
15
20
minutes
1.5
1.0
0.5
1
13
2
BW ID 1
i.s.
0.0
5
10
15
20
minutes
MCounts
2
1.50
1
5
0.00
MCounts
7
4
11
10 12 14
13
3
1.00
0.50
5
10
PW ID 2
i.s.
15
20
minutes
1.50
1.00
0.50
PW ID 1
1
2
0.00
MCounts
8 9
6
4
13
5
10
i.s.
15
20
minutes
1.00
0.50
1
13
2
5
CD UE 2
i.s.
78
0.00
MCounts
5
10
15
20
minutes
1.00
CD EU 1
0.50
1
0.00
2
5
5
7
13
i.s.
15
10
133
20
minutes
The list of 38 pyrolysis products with corresponding retention times, m/z fragments
used for quantification and the total yields are provided in Table 5.2.2. These pyrolysis
products, as described in Conti et al. (2014), were grouped into three thermolabile class
fractions: highly carbonised (charred), weakly carbonised hemi/cellulose and weakly
carbonised lignin (see experimental part). The grouping of pyrolysis products into the
fraction was specified in Section 5.2.2.4.
The quantity of evolved pyrolysis products was expressed in terms of “yield” to give a
rough estimate of the mass fraction that was analysed by GC–MS. Table 5.2.6 shows that
the summed yields varied over three orders of magnitude, spanning from 2,6 106 µg g-1
(PW ID 2, pyrolysis of pine wood at 400 °C) down to 192 µg g
-1
for the biochar
characterized by the highest temperature production and the highest content of ash (corn
digestate biochar). Not surprisingly, the highest yields were obtained with low
temperatures (340 and 400 °C). The biochar synthesized at 600°C and at high ash contents
(corn digestate) gave very weak signals (Figure 5.2.3) because of high thermal stability and
therefore limited the ‘‘pyrolysability’’ of large polyaromatic clusters. Fig. 5.2.3 shows
more details on the yield of biochar as a function of temperature, as well as the charred
content of biochar produced under the different conditions. It can be seen that the yield of
biochar decreases while the BC increases with the pyrolysis temperature in the studied
range.
The influence of feedstock type on the degree of carbonisation is evidenced by results
Py-GC-MS obtained from biochar of corn digestate. This feedstock at 400 °C produced a
thoroughly carbonised biochar with the high charred percentage and the almost complete
disappearance of phenol and methoxyphenols from the pyrolysates (Fig. 5.2.3). This result
could be explained also by the fact that the corn digestate biochar is characterized by the
relatively high content of ash. Moreover, it is interesting to note that biochars more instable
(PW ID 2 and BW ID 2) are those with the higher level of PAHs (PW ID 2) and lower
level (BW ID 2).
In summary, these two techniques provided detailed and consistent information
concerning the chemical characterisation and the stability of biochar samples. For
pyrolysis-GC/MS, the sum of the charred products is a representative parameter of the
relative proportion of BC in biochar. While for HyPy, the stable fraction of biochar can be
defined as the portion of the biochar stable under HyPy conditions (BCHyPy). The degree of
carbonisation, based mainly on the characterisation of the samples using Py-GC-MS and
134
HyPy, increased in the order PWID2 ~ BWID2 < BWID1 ~ CDEU2 < PWID1 ~ CDEU2;
his was corroborated by the elemental analysis. It can also be inferred from the results that,
for a more complete carbonisation of the biochar, it is important the feedstock type and the
final pyrolysis temperature, but also that at high temperature the chemical characteristics of
biochar are influence to a lesser extent by feedstock. In particular, the concentrations of
PAHs ranged between 2.2 (BW ID 2) and 2.9 µg g-1 (PW ID 1), thus always below the
levels recommended by International Biochar Initiative or European Biochar Certificate (412 µg g-1). Therefore, pyrolysis processes will need to be set up to maximise the overall
benefit, not only the yield of stable biochar, and will therefore be very case specific.
Table 5.2.6. Yields in µg g-1 and benzene/toluene ratio (B/T) from Py–GC–MS of
biochars.
Sample Temp.
°C
Py-GC-MS yields
(µg g-1)
Charred Lignin Proteins Holocellulose B/T
%
%
%
%
PW ID 1
600
192
99.5
-
0.5
-
3.2
PW ID 2
400
2654434
36.6
61.9
0.2
1.4
0.49
CD EU 1
600
572
86.6
13.4
-
-
2.6
CD EU 2
400
283
96.1
3.6
0.3
-
6.0
BW ID 1
600
502
99.8
-
0.2
-
8.9
BW ID 2
340
66152
37.4
62.5
0.001
0.001
0.49
5.2.3.4. HyPy and Py-GC-MS: molecular characterization of labile fraction of biochar
As highlighted in the previous section, the stable fraction of biochar is one of the key
parameters to be considered in defining biochar production conditions. However, the labile
fraction, which evolves during its storage in soil, is also very important. This fraction is
highly likely to impact on microbial activity (Lehmann et al., 2011, Ameloot et al., 2013),
and therefore affects the functioning of the soil as a whole, including the balance of
indigenous labile pools (de Graaff et al., 2010, Ameloot et al., 2014). In fact, the
microorganisms can utilize a number of labile biochar constituents as an energy source
(Cross and Sohi, 2011). These are presumably either relatively untransformed biomass
components that have not been subjected to volatilization during pyrolysis (Ronsse et al.,
135
2013) or volatilized compounds that have recondensed in the biochar matrix during
pyrolysis (Imam and Capareda, 2012; Kloss et al., 2012). Moreover, many biochar
associated labile components have biocidal activity (Graber et al., 2010), which may
increase its stability against biotic decomposition.
As well as isolating the BCHyPy fraction, HyPy also allows the characterisation of the
non-BCHyPy material at a molecular level by GC-MS analysis. Fig. 5.2.5 shows mass
chromatograms of the non-BCHyPy fraction derived from biochars, which was found to
contain a high abundance of PAHs. The PAHs detected and quantified (Table 5.2.7) in the
biochars ranged from 2-ring compounds (naphthalene) to 7-ring compounds (coronene),
with the 4-ring compound pyrene being the most abundant. This range of ring size is
consistent with the PAHs distribution found in the non-BCHyPy fraction generated by the
HyPy of 5 archaeological charcoals (Ascough et al., 2010), and the definition of BCHyPy as
being composed of PAHs with >7 rings proposed by Meredith et al. (2012). Their presence
in the non-BCHyPy fraction will be due to their greater volatility relative to the larger more
condensed and refractory aromatic domains which form the BCHyPy.
µg g-1
Figure 5.2.5. Concentrations of PAHs released by the HyPy of biochar. Mean values in µg
g-1 d.w. and standard deviation from two replicates.
9000
8000
7000
6000
5000
4000
3000
2000
1000
0
PAHs HyPy
PWID1 PWID2
BWID1 BWID2
CDEU1 CDEU2
The total mean concentration value of PAHs in biochars (HyPy determination) ranged
between 7531 (PW ID 2) and 43 µg g-1 (CD EU 1) (Table 5.2.7 and Fig. 5.2.5). However,
it is interesting to note that at high temperature also the concentrations of PAHs in labile
136
fraction of biochar determinate by HyPy are influence to a lesser extent by feedstock.
Moreover, the biochars produced at 600 °C, as for the PAHs concentration determined by
soxhlet extraction, have the PAH levels lower (43-1520 µg g-1) than that at low
temperature (920-7531 µg g-1).
Almost all PAHs were detected and quantified in the biochars at lower temperature
(Table 5.2.2), while in all of the biochars at 600 °C, naphthalene, acenaphthylene,
acenaphthene,
methylchrysene,
benzo(b)fluoranthene,
benzo(k)fluoranthene,
benzo(a)pyrene, dibenz(a,h)anthracene, benzo(ghi)perylene, indeno(1,2,3-c,d)pyrene and
coronene were not detected. The individual concentrations of the PAHs in biochars are
presented in Table 5.2.7 and typical distribution profiles are shown in Fig. 5.2.6. A detailed
analysis of the contribution of the individual PAHs in biochars produced at 600 °C
indicated the dominance of phenanthrene (8-32% of the total PAHs), fluoranthene (15-28%
of the total PAHs) and pyrene (36-61% of the total PAHs) in all the samples studied. While
in the wood biochars at lower temperature the PAHs with 5–7 rings composed almost the
majority of PAHs (PW ID 2 = 48%, BW ID 2 = 55%). Therefore, it can be assumed that
biochar generated at a temperature of 340-400 °C will have an aromatic structure that is
not sufficiently condensed to be entirely captured in the analytical window of HyPy.
However, also in the biochar CD EU 2 (400 °C), phenanthrene, fluoranthene and pyrene
dominated the PAH profiles, supplying 28 %, 14 % and 40% of the total PAH
concentrations, respectively. This result could be explained also by the fact that the corn
digestate biochar is characterized by the relatively high content of ash.
The distribution of pyrolysis products is sensitive to the feedstock, as well as
production conditions (Figure 5.2.4 and Table 5.2.8). The pyrolysates of all biochar
samples were featured by the presence of aromatic hydrocarbons including benzene,
benzene derivatives, and polycyclic aromatic hydrocarbons (PAHs; e.g., naphthalene,
phenanthrene). Aromatic hydrocarbons (e.g., benzene, toluene, C2-benzenes, naphthalene,
phenanthrene, diphenyl) along with benzofurans were grouped into a single family of
compounds representing the charred fraction of biochar (C in Table 5.2.5). The high
proportion of these products in the pyrolysates (% charred) which ranged from 36.6% (PW
ID 2) to >99% (PW ID 1 – BW ID 1), indicative of charred biomass. This is in accordance
to Kaal et al. (2009) who proposed that benzene, toluene, naphthalene, diphenyl and
benzofuran could be associated specifically to the charred fraction of BC.
137
In addition, the degree of de-alkylation might be a proxy of thermal alteration (Kaal et
al., 2012). The de-alkylation degree can be estimated in Py–GC–MS from the ratio of
parent/alkylated compound, such as benzene/toluene (B/T), ratios. B/T ratios for the
biochar pyrolysates ranged between 0.49 and 8.9 (Table 5.2.5) and tended to increase with
decreasing overall yields and with increasing the relative abundance of pyrolysis products
indicative of charring (% charred).
The pyrolysis products lignin markers, represented by 2-methoxyphenols (guaiacols),
4-vinylguaiacol,
4-methylguaiacol,
4-ethylguaiacol,
4-methylsyringol
and
2,5-
dimethoxyphenols (syringols), were abundant in the pyrolysate of PW ID 2 and BW ID 2
biochar, while these lignin markers (Ralph and Hatfield, 1991) were not detected in PW ID
1, BW ID 1, CD EU 1 and CD EU 2. The phenols and methylphenols, which are less
specific lignin markers, were abundant in PW ID 2 and BW ID 2, but were detected also in
corn digestate biochars. In this case, the phenols and methylphenols are therefore of little
diagnostic value with respect to highly or weakly pyrolysed lignin.
Figure 5.2.6. Total ion chromatograms from HyPy of biochars PW ID 2 and BW ID 2
showing the PAHs present in the non-BCHyPy fraction.
PW ID 2
Abundance
2e+07
1.6e+07
1.2e+07
8000000
4000000
0
10.00
20.00
40.00
30.00
50.00
60.00
Abundance
BW ID 2
5000000
4000000
3000000
2000000
1000000
10.00
20.00
30.00
40.00
138
50.00
60.00
Table 5.2.7. Observed concentration of PAHs released by the HyPy of biochar samples.
Naphthalene
128
2
PW ID 1
(µg g−1)
n.d.
Biphenyl
154
2
n.d.
96 ± 24
n.d.
n.d.
n.d.
28.0 ± 0.5
Acenaphthene
154
3
n.d.
77 ± 11
n.d.
n.d.
n.d.
47.5 ± 2.2
Fluorene
166
3
3.70 ± 0.7
416 ± 45
n.d.
14.2 ± 0.9
14.4 ± 0.16
161 ± 33.9
Phenanthrene
178
3
186 ± 41.4
1019 ± 32
3.4 ± 0.07
257 ± 46.1
579 ± 9.2
335 ± 80.4
Anthracene
178
3
20.5 ± 0.7
166 ± 9
1.3 ± 0.01
13.4 ± 2.1
18.4 ± 0.3
74 ± 14.7
Fluoranthene
202
4
111 ± 7.3
548 ± 27
11.9 ± 2.8
126 ± 5.2
225 ± 7.1
289 ± 61.1
Pyrene
202
4
239 ± 49.2
938 ± 14
26.0 ± 6.3
406 ± 71.6
540 ± 14.5
432 ± 102
Chrysene
228
4
19.8 ± 1.4
265 ± 24
n.d.
5.87 ± 0.7
53.2 ± 3.5
186 ± 45
Benzo[a]anthracene
228
4
10.7 ± 1.3
346 ± 13
n.d.
18.3 ± 3.3
19.0 ± 0.4
150 ± 26.0
Methylchrysene
242
4
n.d.
311 ± 6
n.d.
10.0 ± 1.5
14.0 ± 0.2
234 ± 22.7
Benzo[b]fluoranthene
252
5
n.d.
734 ± 30
n.d.
21.5 ± 1.0
32.9 ± 0.9
384 ± 59.9
Benzo[k]fluoranthene
252
5
n.d.
470 ± 18
n.d.
17.4 ± 1.8
24.4 ± 1.7
266 ± 58.6
Benzo[a]pyrene
252
5
n.d.
463 ± 26
n.d.
8.1 ± 1.2
n.d.
243 ± 45.0
Indeno[1,2,3-cd]pyrene
276
6
n.d.
601 ± 56
n.d.
4.76 ± 0.7
n.d.
346 ± 28.8
Dibenzo[a,h]anthracene
278
6
n.d.
159 ± 16
n.d.
2.65 ± 0.04
n.d.
102 ± 7.8
Benzo[ghi]perylene
276
6
n.d.
682 ± 86
n.d.
15.8 ± 1.2
n.d.
378 ± 51.8
Coronene
300
7
n.d.
186 ± 16
n.d.
n.d.
n.d.
115 ± 2.1
591 ± 93.7
7531 ± 270
43 ± 9.1
920 ± 131
1520 ± 10.8
3778 ± 593
PAHs
Total PAHs
Mol. Wt.
Rings
PW ID 2
(µg g−1)
57 ± 30
CD EU 1
(µg g−1)
n.d.
CD EU 2
(µg g−1)
n.d.
BW ID 1
(µg g−1)
n.d.
BW ID 2
(µg g−1)
8.0 ± 3.8
139
Table 5.2.8. Concentrations of the pyrolysis products of biochar. Mean values (µg g-1) and
standard deviation from two replicates.
Sample Id.
Compound name
Benzene
Hydroxyacetone
Dimethylfuran
Pyrrole
Toluene
2-Methyltiophene
Furaldehyde
o-Xylene
m-p-Xylene
Styrene
Ethyl-benzene
Phenol
Benzofuran
Benzonitrile
Indole
3-Methylphenol
4-Methylphenol
Guaiacol
Methyl-benzofuran(1)
Methyl-benzofuran(2)
Methyl-benzofuran(3)
2-Ethylphenol
2,5-Dimethylphenol
2,3-Dimethylphenol
3-Ethylphenol
Naphthalene
Catechol
2-Methylnaphthalene
1-Methylnaphthalene
4-Vinylguaiacol
4-Methylguaiacol
Syringol
Biphenyl
4-Ethylguaiacol
4-Methylsyringol
o-Isoeugenol (i.s.)
Fluorene
Phenanthrene
Anthracene
Fluoranthene
Pyrene
BW ID 1
BW ID 2
-1
µg g
pyrolysed
341 ± 83
4592 ± 771
n.d.
n.d.
n.d.
773 ± 23
n.d.
n.d.
38 ± 9.5
9452 ± 1106
n.d.
n.d.
n.d.
136 ± 34
3.9 ± 1.1
909 ± 40
3.7 ± 0.2
3274 ± 385
22.8 ± 1.1
830 ± 34
n.d.
79 ± 5.1
n.d.
15620 ± 2701
n.d.
1345 ± 137
n.d.
n.d.
2.4 ± 0.3
99.0 ± 0.7
n.d.
4340 ± 249
n.d.
7307 ± 92
n.d.
1563 ± 41
n.d.
319 ± 12
n.d.
660 ± 1
n.d.
911 ± 43
n.d.
312 ± 11
n.d.
2910 ± 18
n.d.
2082 ± 13
n.d.
620 ± 22
63 ± 8.7
863 ± 74
n.d.
4149 ± 1047
5.2 ± 0.8
526 ± 53
5.6 ± 0.4
264 ± 29
n.d.
46.1 ± 0.1
n.d.
1001 ± 192
n.d.
n.d.
8.7 ± 1.1
44.5 ± 1.5
n.d.
995 ± 111
n.d.
n.d.
71 ± 1.1
60 ± 3.1
2.4 ± 0.12
94 ± 7.1
2.5 ± 0.05
16.8 ± 0.6
2.4 ± 0.20
12.4 ± 2.0
n.d.
3.8 ± 1.1
n.d.
3.4 ± 1.0
PW ID 1
PW ID 2
-1
µg g
pyrolysed
109 ± 6.8
265527 ± 39092
n.d.
n.d.
n.d.
17.2 ± 1.5
n.d.
30 ± 18
34 ± 5.6
544318 ± 61563
n.d.
n.d.
n.d.
n.d.
6.3 ± 1.4
51.4 ± 4.2
4.8 ± 1.2
183801 ± 16905
7.9 ± 0.7
51.4 ± 1.9
n.d.
n.d.
n.d.
662373 ± 470703
5.5 ± 0.6
79.1 ± 2.5
0.90 ± 0.14
n.d.
0.39 ± 0.02
4.8 ± 0.2
n.d.
279125 ± 86793
n.d.
497598 ± 265920
n.d.
81.3 ± 1.2
0.34 ± 0.08
19.3 ± 1.7
1.1 ± 0.22
18.5 ± 1.6
1.1 ± 0.22
51.7 ± 3.8
n.d.
18.6 ± 7.5
n.d.
220813 ± 88639
n.d.
25.6 ± 10.1
n.d.
27.4 ± 6.3
15.5 ± 0.05
50.4 ± 9.3
n.d.
129 ± 13.1
1.60 ± 0.03
31.9 ± 8.5
1.92 ± 0.07
17.0 ± 5.6
n.d.
2.3 ± 0.2
n.d.
125 ± 31.6
n.d.
n.d.
1.53 ± 0.09
4.1 ± 1.6
n.d.
31.9 ± 11.7
n.d.
n.d.
22 ± 2.3
15.7 ± 4.8
n.d.
3.1 ± 0.5
n.d.
6.2 ± 0.8
n.d.
1.3 ± 0.2
n.d.
0.9 ± 0.1
n.d.
0.7 ± 0.1
140
CD EU 1
CD EU 2
-1
µg g
pyrolysed
237 ± 10
91 ± 16
2
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
91 ± 42
15.3 ± 1.3
n.d.
n.d.
n.d.
n.d.
14.4 ± 1.0 1.18 ± 0.01
15.2 ± 0.3 1.55 ± 0.05
23 ± 5.4
10.7 ± 2.3
n.d.
n.d.
n.d.
n.d.
17 ± 3.0
n.d.
17.6 ± 0.9
35 ± 8.0
n.d.
0.94 ± 0.36
17 ± 13
2.84 ± 0.08
27 ± 26
4.24 ± 0.18
n.d.
n.d.
n.d.
n.d.
n.d.
5.3 ± 1.3
n.d.
6.7 ± 1.8
n.d.
n.d.
11 ± 8.3
0.92 ± 0.11
10 ± 7.5
1.16 ± 0.15
11 ± 7.6
0.93 ± 0.09
50 ± 4.9
76 ± 20
n.d.
n.d.
6.4 ± 2.1
14.1 ± 3.3
6.4 ± 1.7
9.4 ± 1.9
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
9.4 ± 0.8 4.38 ± 0.99
n.d.
n.d.
n.d.
n.d.
61 ± 9.3
74 ± 18
3.9 ± 0.1 0.93 ± 0.11
3.9 ± 1.9 0.25 ± 0.02
n.d.
n.d.
n.d.
n.d.
n.d.
n.d.
5.2.4. Conclusions
The aim of this study was to examine molecular proxies of the thermal stability of the
carbon in biochar produced from different feedstocks and under different production process
and in the context of its carbon sequestration potential. HyPy was applied to isolate the carbon
component most likely stable in the environment on centennial timescales and the Py-GC-MS
providing qualitative and quantitative information on stability of biochar.
The results presented in this study demonstrated that HyPy is a rapid and convenient
technique for the quantification of BC and determination of the stable carbon in biochar. In
support of HyPy method, a strong relationship was found between the H/C and O/C values of
biochar samples and the BCHyPy values found. Moreover, the ability of the HyPy method to
determine BC in the biochar was evaluated comparing the results with Py-GC-MS. The
results showed that these two techniques provided detailed and consistent information
concerning the chemical characterisation and the stability of biochar samples. As well as
isolating the BCHyPy fraction, HyPy also allows the characterisation of the non-BCHyPy
material at a molecular level by GC-MS analysis.
The influence of feedstock type and pyrolysis conditions on the degree of carbonisation and
other biochar properties was evidenced by HyPy and Py-GC-MS. In particular, the fraction of
BC in wood biochar increases with increasing pyrolysis temperature levelling the biochar
characteristics from the feedstock. The levels of solvent extractable PAHs and HyPy evolved
PAHs were higher in biochars produced at the lower temperatures. This means that biochar
exposed to higher pyrolysis temperatures contains a higher proportion of the stable fraction
than biochar produced at low temperatures suggesting that, from a carbon sequestration point
of view, high temperature pyrolysis biochar is preferable. However, studies indicate that from
the point of view of sequestering maximum amount of carbon per unit of feedstock, lowtemperature conversion processes might perform as effectively as higher temperature
pyrolysis processes. Therefore, this aspect requires further investigations.
141
6. Conclusions
The study presented in this thesis was targeted to expand our understanding on the
application of biochar in the environment. In particular, focusing on the potential risks
associated to the application of biochar in soil biochar due to the presence of harmful
substances as well on the stability of biochar.
To the purpose of evaluating the potential risks arising from the occurrence of
polycyclic aromatic hydrocarbons (PAHs) sorbed onto biochar, an analytical method was
developed for the determination of the 16 USEPA-PAHs in the original biochar and soil
containing biochar. The concentration of these PAHs along with the 15 EU-PAHs, priority
hazardous substances in food, was determined in a suite of currently available biochars for
agricultural field applications, which were derived from a variety of parent materials and
pyrolysis conditions. The method consisted in surrogate PAH spiking, prolonged soxhlet
extraction with acetone-cyclohexane, SPE clean-up and GC-MS analysis. The method was
successfully validated with a certified reference material for the soil matrix. In the absence
of commercially available reference materials for charcoal the method could not be fully
validated for biochar. However, the recoveries of surrogate (perdeuterated) PAHs were
satisfactory for almost all of the many investigated biochars from different substrates and
synthesis conditions. The participation to a laboratory exercise within the EU-COST
TD1107 enabled a comparison of the method with methods in use in other laboratories.
All the biochars analyzed in this thesis contained the USEPA, as well as some of the
EU-PAHs at detectable levels ranging from 1.2 to 19 µg g-1. Results have indicated that,
considering an application of 20-60 t biochar ha-1, the degree of PAH contamination will
be dependent on both the presence of background PAHs in soil and the concentrations of
sorbed PAHs on the biochar. Moreover, along with PAH levels determined in other
studies, our data suggested that biochars produced by slow pyrolysis from woody biomass
possessed the lowest level of sorbed PAHs (< 10 µg g-1).
Once spread in the soil, biochar could be a source or a sink of PAHs. The
environmental fate of biochar-associated PAHs is still poorly understood due to the paucity
of long-term in-field studies on this topic. Therefore, it is necessary to improve knowledge
of the role biochar plays in sorbing PAHs and on microbial activity and how this
influences the concentration of PAHs in soil and their persistence in the environment.
The changes in PAH content and distribution was examined in a four year study
following biochar addition in soils in a vineyard (CNR IBIMET). The obtained results
142
showed that the biochar addition determined an increase of the amount of PAHs. However,
the levels of PAHs in the soil remained within the maximum acceptable concentration for a
number of European countries. Moreover, the biochar did not reduce the degradation of
PAHs in the investigated agricultural soil, a conspicuous fraction of PAHs was degraded
bringing the PAH levels close to those of the untretaed soil. The absence of an increasing
concentration trend with time indicated that biochar did not act as a sink of environmental
(e.g. atmospheric) PAHs. Therefore, the impact attributable to PAHs following biochar
application to soil can be minimal.
The four years sampling of vineyard soil performed by CNR-IBIMET was exploited to
study the environmental stability of biochar and its impact on soil organic carbon. In the
literature, several approaches have been proposed to assess biochar stability. Yet, there is
no agreed methodology for determining the long-term stability of biochar. In this research,
the stability of biochar produced from different feedstock and under different production
processes was investigated by analytical pyrolysis (Py-GC-MS) and pyrolysis in the
presence of hydrogen (HyPy). In particular, HyPy was applied to isolate the carbon
component most likely to be stable in the environment on a centennial timescales The
findings of this study showed that biochar amendment significantly influence soil stable
carbon fraction concentration during the incubation period. In particular, the obtained
concentrations of stable carbon fraction in the amended soil are significantly higher than
those in the untreated soil. Obviously, the effect of biochar addition in soils on the level of
stable carbon will depend on the BC in the original biochar. The high stable carbon value
found for biochar (83±3.3%) used in this study suggests that it should be quite recalcitrant
to degradation.
Moreover, the HyPy allowed the characterisation on a molecular level of labile carbon
fraction defined as non-BCHyPy fraction. In addition to a number of PAHs, the non-BCHyPy
fraction was also found to contain a significant abundance of n-alkanes, with a marked
predominance of even-numbered homologues. These compounds are probably derived
from lipids, hydrogenated during HyPy.
The results presented in this study demonstrated that HyPy is a valid technique for
isolating and quantifying stable carbon in soil matrices treated with biochar. Moreover, the
ability of the HyPy method to determine stabile carbon fraction in the biochar was
evaluated comparing the results with flash analytical pyrolysis (Py-GC-MS) on a variety of
biochars. In fact, Py-GC-MS can provide information on the thermal labile fraction of
143
biochar at a molecular level. HyPy and Py-GC-MS were applied to biochars deriving from
three different feedstock (woody, herbaceouse and digestate biomass) at two different
pyrolysis temperatures. HyPy and Py-GC-MS evidenced the influence of feedstock type
and pyrolysis conditions on the degree of carbonisation and other biochar properties. In
particular, the stable fraction in wood biochar increases with increasing pyrolysis
temperature levelling the biochar characteristics from the feedstock. This means that, from
a carbon sequestration point of view, a high temperature pyrolysis biochar is preferable.
In general, the obtained results showed that these two techniques provided detailed and
consistent information concerning the chemical characterisation and the stability of biochar
samples. By isolating the stable fraction, HyPy also allowed the characterisation of the
labile carbon fraction at a molecular level by GC-MS analysis.
Biochar has potential as soil amendment for improving soil quality, decreasing
fertilizers losses and store carbon into the soil. Nevertheless, as soil additive, the absence
of phytotoxicity is the minimal requirement. We have showed above that the concentration
of PAHs in biochars from slow pyrolysis of lignocellulosic feedstock is generally
sufficiently low to keep the degree of contamination in soils at safe levels. However,
biochars from sources other than woody or herbaceous biomass could exhibit detrimental
effects on plants. In this context, biochar from poultry litter was investigated in this thesis.
Biochars were prepared by intermediate pyrolysis at different temperatures and compared
with biochars from corn stalk prepared under the same pyrolysis conditions. The
phytotoxicity of these biochars was estimated by means of seed germination tests on cress
(Lepidium sativum L.).
Results obtained show that biochar from poultry litter may exert negative effect at least
at the relatively high level of soil amendment (40 t ha-1). The role of PAHs in the inhibition
of seed germination was excluded. Instead, potential candidates of toxicity were identified
in water soluble and biodegradable components, probably derived from the thermal
decomposition of proteins and lipids. In supporting this hypothesis, the toxicity was
drastically reduced by water extraction or mixing with biologically active materials, while
the water extracts inhibited the germination. Therefore, biochar is not an “intrinsically
safe” material, and every biochar from different processes and/or feedstock has to be
evaluated, checked and possibly treated before the agronomic application.
144
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Acknowledgments
First of all, I would like to express my sincere gratitude to my supervisor, Prof. Daniele
Fabbri. His invaluable support in my research and constant encouragement during all my
PhD have been essential for the completion of this thesis.
Secondly, my acknowledgements to Prof. Colin E. Snape (University of Nottingham) and
to Dr. Will Meredith (University of Nottingham) for hosting me and supervising my work,
organizing the hydropyrolysis experiments of biochar and soil.
Surely, I could not forget Dr. Silvia Baronti (Istituto di Biometeorologia – CNR, Firenze)
for soil and biochar amended soil used for analysis described in Chapters 3.2 and 5.1 and
Dr. Alba Dieguez-Alonso (Technische Universität Berlin) for biochars used for analysis
described in Chapter 5.2. I am grateful to Dr. H.-P. Schmidt who coordinated the WP1 of
the EU-COST TD1107 for the participation to the ring trial. Part of the activity was
conducted in collaboration with the Unità Operativa Biomasse within the framework of the
APQ Ricerca Intervento a “Sostegno dello sviluppo dei Laboratoridi ricerca nei campi
della nautica e dell’energia per il Tecnopolo di Ravenna” “Energia, parte Biomasse”
between Università di Bologna and Regione Emilia Romagna (Italy). The biochars used in
the characterizations reported in Chapter 3.1 are part of the USDA-ARS Biochar and
Pyrolysis Initiative and USDA-ARS GRACENet (Greenhouse Gas Reduction through
Agricultural Carbon Enhancement Network) Programs.
I would still like to acknowledge all professors and colleagues which contributed to the
work presented in this thesis: Prof. Kurt Spokas (USDA) for biochars collection and
characterization, Dr. Cristian Torri (University of Bologna, Department of Chemistry G.
Ciamician) for his work in biochar characterization and germination test, Dr. Roberto
Conti (University of Bologna, Fraunhofer UMSICHT) for synthesis and characterization of
biochar, Dr. Michele Ghidotti (University of Bologna, CIRI EA) Dr. for SPME analysis,
Dr. Dennis Zannoni for ammonium analysis (University of Bologna, CIRSA) and Dr.
Giovanni Marisi for some of the experiments of section 4.
During my PhD, I had the privilege of meeting and collaborating with many precious
people, who helped and supported me in different ways. No words to truly express my
gratitude to Dr. Chiara Samorì (University of Bologna, CIRSA), Dr. Danilo Malferrari
(University of Bologna, CIRSA) and Dr. Chiara Lorenzetti (University of Bologna,
CIRSA).
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Finally, I would like to deeply thank my wife Mariarita, whose unconditional and endless
love has always been a propellant energy for all my achievements.
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