null  User manual
Arctic LTER Streams Protocol
Field and Lab Methods
Toolik Field Station and
RESL at the University of Vermont
Elissa Schuett and Joshua Beneš
Updated May 2013
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Arctic LTER Streams Protocol
Table of Contents
I.
General Toolik Field Sampling Info ................................................................ 3
A. Wet Lab ............................................................................................................... 3
B. Field Sites ............................................................................................................ 4
C. Nutrients .............................................................................................................. 6
D. Rock Scrubs ...................................................................................................... 10
E. Discharge ........................................................................................................... 12
F. TSS .................................................................................................................... 17
G. Dripper ............................................................................................................... 18
H. Macroinvertebrates ............................................................................................ 19
I.
Moss point transects .......................................................................................... 19
J. Fish .................................................................................................................... 20
II. Lab Methods at Toolik ........................................................................................... 23
A. Soluble Reactive Phosphorus ............................................................................ 23
B. Ammonium using the OPA method .................................................................... 27
C. Chlorophyll a ...................................................................................................... 30
III. Lab Methods at the Rubenstein Ecosystem Science Laboratory (RESL) ............... 33
A. Nitrate ................................................................................................................ 33
B. TDP ................................................................................................................... 36
C. TDN/DOC .......................................................................................................... 40
D. Particulate Phosphorus ...................................................................................... 43
E. CHN ................................................................................................................... 45
F. Anions ................................................................................................................ 49
G. Cations .............................................................................................................. 51
H. Alkalinity ............................................................................................................ 55
IV. Appendix A – Formerly Used Methods .................................................................. 59
A.
B.
C.
D.
2x2 rock scrubs.................................................................................................. 59
Epilithic primary productivity ............................................................................... 60
Bioassays of epilithic algae ................................................................................ 61
Ammonium – Phenol method ............................................................................. 63
Appendix B – Locations of Equipment ................................................................... 70
A.
B.
C.
At Toolik............................................................................................................. 70
At the RESL - 2013 ............................................................................................ 71
Stored at the MBL .............................................................................................. 72
V.
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Arctic LTER Streams Protocol
I.
General Toolik Field Sampling Info
Sampling of many different things occurs throughout the summer for the LTER
Streams group. The Kuparuk River and Oksrukuyik Creek are the main sampling
locations for the LTER, but other sites may be added in different years. These
protocols are specific to the long-term sampling of the Kuparuk River near the
Haul Road and Oksrukuyik Creek near the Haul Road, and Imnavait Creek
surrounding the access road and thermokarst. Additional streams that are
sampled are not discussed in detail because the streams change regularly and
sampling protocols may differ slightly, however, all chemistry methods remain the
same, or allow for overlap of analyses. Other types of streams that have been
studied have included streams that are impacted by thermokarst, tributaries of a
stream network, and spring, glacier, and mountain streams. The Kuparuk has
had phosphorus added to it every year since 1983, Oksrukuyik had phosphorus
and nitrogen added 1991-96. Reference, recovery, and fertilized reaches are
included in the sampling regime. Each stream is sampled three times during the
summer for nutrients and benthic chlorophyll (rock scrubs). Regular discharge
measurements are taken on both streams. Arctic grayling are caught and
released, both adults and young of the year. Macroinvertebrates, both with
Surber samplers and drift nets, are collected at least twice from the Kuparuk, less
frequently from Oksrukuyik. Moss point transects are done on the Kuparuk twice
during the field season.
A. Wet Lab
The Wet Lab at Toolik is a highly used lab space – many people are often
coming and going. It is important to follow some rules so that everything runs
smoothly – including both personal safety and good science.
1. Some rules
a. Outside shoes are not allowed
b. Designated, close toed lab shoes are required
c. Stay out of the RAD area
d. Follow the Nutrient RA’s rules and guidelines
e. Sign up for analyses
2. Acid washing
a. Two acid baths are on the front porch – one for NH4 equipment and
one for phosphorus/anything else – only use the NH4 bath if the
Nutrient RA says it is ok
b. rinse everything that is going in the bath with DI 3x
c. remove all tape!!! It makes everyone happier – when tape has been
acid washed it becomes very gooey, just making it more difficult to
remove the next time that you try.
d. using long rubber gloves, rubber apron, and goggles put lab ware into
the bath
e. let soak for ~1hour
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Arctic LTER Streams Protocol
f.
g.
h.
i.
• only soak filter cartridges with metal for ~5minutes!
remove the items from the bath – wearing the gloves/apron/goggles
rinse with DI 3X – pour the waste DI/acid into a bucket to be disposed
of in the acid waste (or to mop the floor with)
let dry
put items away
B. Field Sites
1. Kuparuk River
a. The original, 1983 phosphorus dripper was located at what is called
0.0k – all other stations are named as a distance from that location.
Positive numbers are downstream; negative numbers are upstream.
Stations downstream of the dripper were in the fertilized reach,
upstream are reference stations.
b. The phosphorus dripper was moved downstream in 1985 to the
location called 0.56k. It remained at that location until 1995.
c. In 1996 the dripper was moved downstream again to 1.4k and
remained there until 2010. Stations upstream of the dripper were in a
recovery reach. Downstream of the current dripper location is fertilized
and upstream of the original dripper is still reference.
d. In 2011, a second dripper was added at the 0.0k location. Phosphoric
acid is added at a rate of 2.4ml/min. This is in addition to the dripper
which has been at 1.4k since 1996, dripping at 1.2ml/min. The area
between 0.0k and 1.4k has become known as the re-fertilized zone.
This zone is broken up into two “Re-fert” zones, divided at the location
of the dripper at 0.56k from 1985-1995.
e. Stations for long-term data remain the same from year to year, but
some stations are sampled for different things, depending on habitat
characteristics. Most stations that are frequented are riffles.
Station Name
-0.7k
-0.47k
-0.377k
Reach type
Reference
Reference
Reference
-0.177k
Reference
0.0k
Reference
0.3k
Reference
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Arctic LTER Streams Protocol
Samples Collected
YOY
YOY
Nutrients, rock scrubs,
macroinvertebrates, moss
point transect
Nutrients, rock scrubs,
macroinvertebrates, moss
point transect
Original 1983 phosphorus
dripper, YOY, moss point
transect, macroinvertebrates
YOY, moss point transect
0.5k
Reference
0.56k
Reference
0.59k
Fertilized/Recovery
0.6k
0.74k
0.85k
1.0k
Fertilized/Recovery
Fertilized/Recovery
Fertilized/Recovery
Fertilized/Recovery
1.15k
1.2k
1.39k
1.4k
Fertilized/Recovery
Fertilized/Recovery
Fertilized/Recovery
Fertilized
1.8k
Fertilized
2.0k
Fertilized
2.5k
Fertilized
3.0k
Fertilized
4.0k
4.1k
Hershey Creek
Fertilized
Fertilized
Tributary
High flow discharge
measurement, fish weir
Nutrients, rock scrubs, moss
point transect
1985-1995 phosphorus
dripper
Discharge measurement
Nutrients and rock scrubs
YOY
Nutrients, rock scrubs, YOY,
and moss point transect
Moss point transect
Moss point transect
Nutrients and rock scrubs
1996-2010 phosphorus
dripper
Nutrients, rock scrubs, YOY,
moss point transects
Nutrients, rock scrubs, moss
point transects,
macroinvertebrates
Nutrients, rock scrubs,
macroinvertebrates
Nutrients, rock scrubs, YOY,
moss point transects,
macroinvertebrates
Nutrients, rock scrubs
Moss point transects
Nutrients
2. Oksrukuyik Creek
a. Oksrukuyik Creek was sampled beginning in 1989 to study baseline
conditions. Fertilization using phosphoric acid was begun in 1991,
increasing SRP by 0.32 uM. Phosphorus addition continued through
1996.
b. Ammonium sulfate was added 1993-1996, increasing N by 7.1 uM.
c. Recovery from nutrient additions was begun in 1997. No further
additions have been made since then.
d. Sampling in the stream occurs primarily at the reference reach stations
to continue baseline monitoring.
e. Discharge was formerly measured downstream of the Haul Road near
the USGS stilling well, however, in 2008 the USGS equipment was
removed due to impending road construction. The discharge
measurement site was moved upstream of the road ~100m.
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Arctic LTER Streams Protocol
Station Name
-0.7k
Reach Type
Reference
-0.3k
Reference
-0.2k
-0.1k
Reference
Reference
0.227k
0.45k
0.482k
0.85k
1.056k
1.370k
Fertilized/Recovery
Fertilized/Recovery
Fertilized/Recovery
Fertilized/Recovery
Fertilized/Recovery
Fertilized/Recovery
1.700k
2.497k
Fertilized/Recovery
Fertilized/Recovery
Samples Collected
Nutrients, rock scrubs,
YOY
Nutrients, rock scrubs,
YOY
YOY
Nutrients, rock scrubs,
YOY
Nutrients, rock scrubs
YOY
Nutrients, rock scrubs
YOY
Nutrients, rock scrubs
Nutrients, rock scrubs,
YOY
Nutrients, rock scrubs
Nutrients, rock scrubs
3. Imnavait Creek
a. Imnavait Creek is sampled at least twice during each summer by the
streams group in the thermokarst-impacted reach.
b. Nutrient samples and total suspended sediments are collected at each
station.
c. Filtering is often done in the lab because more particulate matter is in
the water, making syringe filtering in the field difficult.
d. It is important to collect from areas of running water, which can
sometimes be difficult in dry years
Station
Name
A
AA
B
C
Station Type
Station Description/Location
Reference
Reference
Impacted
Impacted
D
E
Impacted
Downstream of
impacted area
Upstream of the boardwalk
Inlet to 2nd pool upstream of the culvert
Outlet of pool ~50m below access road
~250m downstream of road (wooden stake
and rebar mark the station)
50m upstream of the black diamond sign
~50m upstream of Haul Road – inlet to the
large pool
C. Nutrients
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Arctic LTER Streams Protocol
On a nutrients day water samples are collected for NH4 and SRP to be run in Wet
Lab at Toolik. Sestonic chlorophyll a samples are also collected and run at
Toolik. Other samples are collected, but preserved for analyses at the RESL or
other home institutions, including NO3, TDP, TDN/DOC, PP, PNPC, anions,
cations, and alkalinity. In addition to water sample collection, pH, temperature,
and conductivity are also measured.
1. Pre-field prep – what to take
Pack everything the day before going into the field – it makes everyone’s life
easier
a. Field book and pencils
b. amber 1 L bottle for chlorophyll a samples
c. Station bags containing:
Item
Analyses
125 mL HDPE amber
NH4/SRP (reuse, must
be acid washed/DI rinsed
between sampling)
(1/regular, 2/rep station)
Nitrate(1/regular, 2/rep
station)
Cations (only done at rep
station)
TDN/DOC (1/regular,
2/rep station), TDP
(1/regular, 2/rep station),
Alk (only done at rep
stations)
Anions (only done at rep
stations)
PP, PCN (1 each/station)
PP, PCN (1 each/station)
Cations (only done at rep
station)
Chl a (1/station)
60 mL LDPE
60 mL HDPE square
60 mL HDPE narrow
mouth
30 mL HDPE
47mm Petri dish
25 mm filter cartridges
Whatman 45um Puradisc
filters
10ml Falcon Tube
Total needed per
station
1/regular station, 2/rep
station
1/regular station, 2/rep
station
1/rep station, 0 per
regular
2/regular, 5/rep
1/rep station, 0/regular
2
2
1/rep station
1
d. 25 mm filter cartridges can be pre-loaded with ashed 25mm GF/F the
morning of field collection (to pack cartridges always use forceps,
never touch the filter! Place in the center of the metal disc, place the Oring on top of the filter, screw the top part of the cartridge on tightly, but
don’t over-tighten or the filter might break).
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Arctic LTER Streams Protocol
e. Syringe box – at least 2 140cc clean syringes, filter forceps, sharpie
f. Extras bag - Forceps, extra filters, an extra filter holder and O-ring,
labeling tape and syringe, sharpie, batteries
g. Calibrated pH/conductivity meter and probes
• set up and calibration - use the pH buffers in Wet Lab to calibrate
the Hach pH probe before use each day
o use pH 7 and 10
o rinse with DI between each buffer
o make sure the electrode has gel – when it is cold this can get too
thick so you might need to try to shake it or click more gel if
using the electrode with gel replacement capsules
o follow the instructions in the manual for calibration
o pour waste into the pH buffer waste container
• make sure there is extra buffer in the case so that field calibration
can be done if necessary
2. Field methods – what to do
a. Nutrients, sestonic particulates, and ions
•
At each station, go to the center of the stream and rinse the syringe
3x with stream water.
•
Fill the syringe with stream water, mount the 25mm filter cassette
onto the syringe and push 5-10 ml of water through the filter to
rinse – also check to make sure no water is coming out of the sides
of the cartridge – the filter might not be seated correctly if you see
water. When filtering always point the syringe towards the ground.
Fill bottles with filtered water, rinsing each sample bottle 2x with
filtered stream water before filling. Keep track of the volume
filtered.
•
After the last water has gone through the filter push a small amount
of air through the filter by removing the cartridge from the syringe
(or turning off the stop-cock) pulling a small amount of air in the
syringe and then pushing it through the filter (if you don’t remove
the cartridge you will pull air through, possibly breaking the filter),
remove the filter with the forceps and place it in a petri dish labeled
for particulates (see below, #8).
o Cations – use a .45um filter for the cation sample – sometimes it
can be very difficult to use these filters so a caulk gun can be
helpful, but be careful to not break the syringe! The square 60mL
bottles are used for cations so that you remember to use the
other filter – so if you run out of squares, it is ok to just use a
regular 60ml.
•
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Record the volume of water filtered through the filter directly
on the labeled petri dish with a sharpie.
Arctic LTER Streams Protocol
•
Avoid making any physical contact with the filtered water!
Your skin can contaminate the water for certain analyses. Keep
mosquitoes and unfiltered stream water out of the sample bottles.
b. Sestonic Particulates (PN, PC, PP)
•
After each filter has had at least 500 mL (for Kup and Oks, you may
not be able to filter 500 mL on other streams) of water run through
it, remove the filter with the forceps and place it into one of the Petri
dishes (it doesn’t matter which one). The dishes should be prelabeled with stream, station date, and either PN/PC (particulate
nitrogen and particulate carbon), or PP (particulate phosphorus).
Place the other filter into the second pre-labeled Petri dish.
IMPORTANT: record the volume of water filtered through the
filter directly on the labeled Petri dish with a sharpie!!
•
Remember to dry field filter blanks for each type of particulate
sample (1/transect). The blanks should be from the same box as
the sample filters and should be dried on the same day.
c. Sestonic chlorophyll a concentration
• At each station, rinse the syringe 3x with stream water. Take the
samples from the center of the river.
• Fill the syringe with stream water, then attach the 47mm filter
cassette (with filter) to the syringe and push water through the filter.
Repeat this step until 500 mL of stream water has passed through
the filter.
• Push a small amount of air through the filter cassette using the
syringe (this pushes the remaining ~5mL of water through the filter
and prevents sample loss in the transfer)
• Remove the filter from the cassette and place in a falcon tube, and
pace this tube in a amber dark bottle so that the Chlorophyll levels
don’t change.
d. pH/conductivity
• put the electrode directly in the stream, but in an area that is not too
turbulent
• if the pH is reading strangely try recalibrating in the field with buffer
that is kept in the case
o if still having problems, try reading the pH in a sample bottle
instead of directly in the water
o check the amount of gel on the electrode
o if all else fails, read the samples back in the lab with a different
pH probe
3. Return from the field – what to do with all of the samples you just
collected
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Arctic LTER Streams Protocol
a. Rinse syringes and filter cartridges with DI 3X – let dry in clean bin (if
you don’t rinse these before they dry you must acid wash them)
• Remember NOT to acid wash the chlorophyll syringes or filter
cassettes between uses; acid rapidly degrades chlorophyll, so
traces of acid from the acid bath could affect the chlorophyll in the
samples.
b. Open pH/conductivity meter box to let air dry
c. Some samples are run immediately, and others are preserved for later
analysis. The following chart tells you where everything goes from a
nutrients day.
Sample
NH4/SRP
TDP
TDN/DOC
Cations
Nitrate
Alks
Anions
Sestonic particulate phosphorus filter
Sestonic particulate carbon and
nitrogen filter
Sestonic chlorophyll a
What happens to it
Run samples on day collected, if can’t
be run w/in 24hrs, freeze
Acidify w/ 100uL 6N HCl, refrigerate
Acidify w/ 100uL 6N HCl, refrigerate
Acidify w/ 100uL 6N HCl, refrigerate
Freeze
Refrigerate
Refrigerate
Dry in oven 60oC for 24 hours, then
store for shipment
Dry in oven 60oC for 24 hours, then
store for shipment
Put in designated 15cc centrifuge tube,
freeze 1 hour – 1 week, analyze on
fluorometer (see methods in lab
analysis section)
D. Rock Scrubs
Algae growing on the stream bottom rocks are an important component of
primary productivity. Therefore, samples of epilithic algae are scrubbed off of the
rocks and their chlorophyll content is measured. Benthic particulate nutrients are
measured as well.
1. Pre-field prep – what to take
a. field book and pencil
b. wash basin
c. small steel bristle scrub brush
d. 500-ml wash bottle (whole rock method)
e. 250 mL sample bottles – two for each station, reuse dedicated bottles
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Arctic LTER Streams Protocol
f.
g.
h.
i.
2x2 slide holders (2x2 method)
3-5 60 ml bottles or 56 ml centrifuge tubes. (2x2 method)
funnel (optional)
black garbage bags
2. Field methods – what to do
Whole rock scrubs - The 2 x 2 rock scrubs outlined in the Appendix is a fast
method used to determine the chlorophyll α and particulate nutrients in the
epilithic algae. Whole rock scrubs are a more thorough method for the same
purpose. Once a summer, whole rock scrubs were conducted at the same time
as 2 x 2 scrubs to compare the results and efficiencies of the two methods. Note:
Since about 2004, we’ve been doing whole rock scrubs only and have
stopped doing 2x2 since whole rock is the preferred method (Bruce
Peterson; personal communication). The methods below are those of Bruce
Peterson (personal communication) and have been modified into outline form.
a. At each station where whole rock scrubs are to be done, rinse the
wash basin, graduated cylinder, brush, and squirt bottle 3x with river
water.
b. Select several rocks from each station that fit the following criteria
(make careful notes of any rocks that do not fit the criteria):
•
no obvious filamentous algae or moss (to avoid overestimates of
chl due to filamentous algae or moss)
•
fairly smooth surface (uneven surfaces prevent efficient removal of
epilithon)
•
similar size (the rocks should be of appropriate size to form a single
layer of rocks that fills the bottom of the wash basin).
c. Remove the rocks from the basin and gently set them aside, right-side
up. Place them in shallow water if you are able to keep track of which
rocks you chose
d. Fill the 500ml rinse bottle with river water. You will need to use 2-500
ml rinse bottles of water to have a 1L initial volume. All of this water will
be needed, so do not spill or waste any.
e. One by one, scrub each rock vigorously and thoroughly with the wire
scrub brush. All scrubbate should fall into the basin.
f. Rinse each rock with the water from the wash bottle, making sure that
all rinse lands in the basin.
g. After all scrubbing is complete, the basin will contain a slurry of 1 liter
(fill the 500ml rinse bottle twice) of water plus all of the scrubbate.
• Stir up the slurry in the basin so that it is homogenous; then fill the
pre-labeled 250-ml bottle with slurry. This is a subsample for
laboratory analysis.
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Arctic LTER Streams Protocol
•
•
Pour out the remaining slurry. Store the bottle for return to camp.
Repeat this procedure twice, so that at each of the stations, you
scrub a total of two basins of rocks.
3. Return from field – what to do with all the samples you just collected
a. Using forceps place pre-combusted Whatman GF/F 25 mm glass fiber
filter onto frit; attach upper reservoir to filter holder.
b. Pipette 5 ml (note if otherwise) from each replicate sample bottle
(shake bottle first) into filter reservoir. Turn on vacuum pump, be sure
not to allow pressure differential to exceed .3 ATMs, to minimize
damage to delicate organisms and don’t let filter pull dry (slowly
release vacuum as final volume of water is filtered).
c. Place filter into centrifuge tubes (Falcon Tubes) and place in the
freezer for at least an hour (no longer than one week). Then add 10ml
of chilled 90% acetone (1mg MgCO3/L). Make sure filter is completely
immersed in acetone, cap tightly and place in light tight box while
processing, then move to cooler with ice for extraction. These filters
will be analyzed for Chlorophyll a content after a 16-18 hour extraction
period. Tubes should be inverted/mixed at least once during extraction,
but allow sufficient time (>4hours) for particles to settle before reading.
d. Once you have filtered 5 ml for chlorophyll for each replicate from a
station, combine the remaining sample (5 for 2x2's and 2-3 for whole
rocks) into a large bottle with lid. Shake the bottle and then filter 5ml
(write volume on Petri dish) of this homogenate through 2 more GF/F
25mm filters for PNPC and PP samples.
e. Place these PNPC and PP filters in petri dishes and place in the drying
oven for at least 24hrs with the lids ajar. (That's it for the PNPCPPstore in a ziploc bag and ship to RESL).see LTER Streams sampling
protocol.
f. Pipette 20 ml of the homogenate into glass scint vials for algal comps.
Preserve with 0.1ml 50% glutaraldehyde. Ship to UVM Rubenstein
Ecosystem Science Lab..
E. Discharge
1. In each stream, discharge will be measured manually at least 6 times
(preferably 8 times) per summer at intervals of approximately one week or
following significant changes in stream levels. The purpose of these
measurements is to create a water level/discharge curve so that the
datalogger water level data can be converted to discharge data.
2. Read and record the river depth on the staff gauge on the side of the
stilling well. The units are in hundredths of feet. Record the time of day
as well.
3. At the designated station in each stream extend a meter tape,
perpendicular to the flow, across the river and secure it on both banks.
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Arctic LTER Streams Protocol
The tape should be relatively taut. This will serve as a reference for doing
the transect.
4. Measure the width of the stream and divide the width into 20 increments.
Write these increments in one column of the notebook.
5. Beginning at one bank, measure and record the depth of the river, then
measure and record the current velocity using the Gurley, MarshMcBirney, or SonTek ADV current meter.
a. If using the SonTek ADV name the file with the first letter of the river (K
for Kuparuk) and the date using DDMMYY format. With the 3D ADV
probe attached to the wading rod, point the probe upstream, stand
downstream of the probe so that you don’t create an eddy. However, if
the water is very clear, you may get a poor reading, in which case one
person should stand upstream of the probe and kick sediment up so
that particles can be read by the probe. Make sure that the person is
far enough upstream so that an eddy is not created in the wake of the
person.
b. Using the top-setting wading rod, set the probe at 60% of stream
depth, i.e., 60% of the way from the surface to the bottom. The wading
rod is in meters and uses a Vernier scale. Each single bar on the rod
indicates 2cm; the double bar is 10cm, and the triple bar is 50cm. So,
to set the wading rod, for example, if the depth is at the mark above
the triple bar the depth is 52cm, you then depress the lever to move
the mobile rod to line the number 5 up with the number 2 on the
Vernier scale at the top. The wading rod is now set at 60% of 52cm, or
31.2cm from the water surface.To begin measuring, set the location
and depth on the SonTek datalogger at the starting edge. Once the
edge is defined, you will need to move away from the edge and any
interference created by the bank.
1. Set the first level of measurement at a location away from the edge
of the bank. Set the depth on the datalogger and adjust the wading
rod.
2. Move to the next interval, there should be about 20 increments
along the width of the stream. Set the location and depth.
o After two intervals, the SonTek should begin to “know” your
interval, however, always check to make sure that it is right. You
will always have to set the depth. It works better to move from
low to high along the meter tape. Approve and warnings or
repeat the measurement – depending on the warning and your
judgment.
c. Move along the tape to the next increment and repeat the depth and
current measurements. Continue until the entire river breadth has been
measured. (If the stream bottom is very inconsistent, you may have to
take measurements at greater or less than the designated increment
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Arctic LTER Streams Protocol
along the tape; be sure to record these discrepancies in the notebook).
Remove the tape after the transect is complete. Read the water depth
again off of the staff gauge at the stilling well. The average of the
depths before and after the discharge measurement is considered the
river level for the given discharge.
d. The discharge is the sum of the products of each individual
measurement (an individual measurement is the water velocity at a
given increment times the depth at that increment times the distance
from the previous increment). This can be computed easily in a
spreadsheet.
e. If the SonTek was used, upon returning to the lab plug the SonTek into
the laptop with the 9pin RS232 cable. Open SonTek FlowTracker
software and connect to the SonTek on Comm Port 1. Save the file
into the appropriate folder.
6. Water level will be recorded continuously in three ways: 1) with a
pressure-sensing probe hooked up to a Campbell Scientific CR-10
datalogger, 2) with a Stevens float-and-pulley water level chart recorder,
and 3) with an Onset HOBO water level logger.
a. The CR-10 datalogger should have 8 fresh D batteries installed at the
beginning of every summer; these should last for the entire summer. A
CR-10 was deployed in Oksrukuyik Creek just downstream of the
Dalton Highway until 2008. Beginning in 2009 the CR-10 was moved
to upstream of the Dalton Highway due to road construction. Doug
Kane’s group (UA-Fairbanks) will deploy a CR-10 in the stilling well at
the Kuparuk.
1. Program the CR-10 datalogger for each parameter using the
owner’s manuals. FIRST MAKE SURE THE TIME IS SET
PROPERLY ON THE DATALOGGER. The LTER computer has a
directory called PC208w. Use the program EDLOG for program
writing and editing. Transfer the program to the CR-10 keyboard
and which can then be downloaded to the datalogger. The CR-10
Prompt Sheet is vital for using the CR10 keyboard and should be
regularly consulted. Decide with Bruce Peterson the number of
times per day the datalogger should record river depth. Consult the
manual for each probe on how to wire the probes to the datalogger.
2. Deploy the probes as soon as possible after arriving at Toolik.
3. Doug Kane’s group (UAF) will deploy the Kuparuk probes.
4. At Oks Creek, place the datalogger inside the yellow weatherproof
metal case with the hole in the bottom; the probes and their cables
should be run through the hole. Using plenty of cable ties, mount
the metal case on the wooden frame supported by rebar (about 5 m
downstream from the stilling well). Drive a piece of rebar into the
14 |
Arctic LTER Streams Protocol
stream bottom in at least waist-high water. Loosely attach a cable
tie to the probe cables, loop the tie around the rebar, and slide the
probes down so that they rest on the bottom. Place some large
rocks on the cables so that the probes stay on the bottom and are
somewhat protected.
5. Data will be downloaded into a storage module for transfer to the
LTER computer at the Toolik Camp (both as an ASCII file and on a
spreadsheet). You must bring the CR-10 keyboard to do this. Data
should be downloaded and backed up weekly, especially in times of
potential flooding. The datalogger should be removed if flooding is
imminent.
6. Back at camp, use the 9-pin SC532 interface cable attached to the
RS232 interface module to communicate between the storage
module and computer. Use the SMCOM program (in PC208
directory) from DOS prompt to dump data. Select COM2 as the
interface port. Select U for uncollected data and C for comma
delineated file. This creates a *.DAT file which can then be
imported into a spreadsheet. Keep the *.DAT files as backups.
Place the *.DAT files in the directory set up for the specific site.
Use the following template for naming downloaded files:
YYCJULFL.dat where YY=year, C = site code, JUL is Julian day
on which downloaded, and FL = file list; for example, 94N22401.dat
is the datalogger file downloaded on Julian Day 224 for the new
reach (Blueberry Cr.) in 1994.
b. A Stevens recorder is deployed in a stilling well upstream of the
pipeline in the Kuparuk. The USGS had a Stevens recorder in a stilling
well downstream of the Dalton Highway crossing at Oksrukuyik Creek
until spring 2008 when it was removed due to potential road
construction.
c. Onset HOBO water level loggers will be deployed in Oksrukuyik and
Kuparuk. A barometric pressure logger should be left logging at the
bank of the Kuparuk River near the water level Onset HOBO or in the
Wet Lab at Toolik Field Station. These loggers will be overwintered to
determine freeze and thaw dates. The first year of use was the winter
of 2008-09. The HOBOs fit inside pipes with end caps that have holes
drilled to allow contact with the water. The pipes are then fitted to
conduit that can fit over rebar that is pounded into the river bottom.
Leashes should also be fitted to secure the HOBO to the bank to
insure they do not get broken away during peak snow melt and ice
flows. A staff gage is also attached to the conduit to determine stage
height.
1. The HOBO should be “calibrated” at the beginning of each season.
Launch the HOBO using HOBOWare Pro – be sure the time is set
15 |
Arctic LTER Streams Protocol
exactly to GMT. Partially fill a bucket of water with lake water. Place
the HOBO in the water. Measure and record the depth of water. Let
the HOBO equilibrate for 15 minutes. Add more water, noting the
time. Measure the depth of water again and let equilibrate. Offload
the data and determine that the time and depth of water is the
same in the HOBO as what you manually recorded.
2. Using the HOBO requires the HOBO shuttle and a computer with
the HOBOWare software. To launch the HOBO connect the HOBO
to the shuttle and the shuttle to the computer. Open HOBOWare
and click launch device. A time delay can be set for when logging
should begin so that all loggers can begin at the same time – this
makes data crunching easier. The time should be set exactly to
GMT. For summer deployment the HOBO should log every 10
minutes. For winter deployment logging should occur once an hour
so that the battery lasts until it can be retrieved in the summer.
3. To upload data from the logger in the field use the shuttle –
unscrew the black cap and line up the flat end of the logger with the
arrow on the shuttle and press the lever. Once the light on the
shuttle is green the data has been uploaded. The logger can then
be put back in the stream to continue logging.
4. To download the data from the shuttle, connect it to the computer
using the USB cable. In HOBOWare click Readout Device. All of
the data will be collected from the shuttle into files for each HOBO.
Each file can then be renamed and saved into an appropriate
folder. The data will be cleared off the shuttle. File saving and
management is important here!
7. Stream height should also be recorded as often as possible by reading the
river depth on the staff gauge and recording it, along with the time (AST)
and date, in the notebook stored in the stilling well (and on the chart
recorder in the stilling well box; write the time, date, and river level on the
recorder paper and draw an arrow to the spot at which the chart recorder
needle is currently located). These data are used to calibrate the manual
discharge measurements with the river height data from the datalogger.
8. At the end of the season, construct a discharge curve for the stream.
a. Convert the stage heights from the manual discharge measurements
(i.e., the measurements taken in part A of this section) from feet (the
measurements taken from the staff gauge) to meters (the depth
measurements recorded by the datalogger at the same time and date
as the manual discharge measurements).
16 |
Arctic LTER Streams Protocol
b. In Excel (5.0 or higher), plot the 6 (or more) discharge measurements
from part A of this section: stage height in meters on the x-axis,
discharge in cubic meters per second on the y-axis.
c. Use the trendline function to draw the line that best fits the discharge
data. You should select for a power curve and select for the equation
and r2 value to be displayed. (Hint: Be sure to display the numbers in
the equation to several decimal places so that your computations in the
next steps are precise.)
d. The equation produced by the relationship will probably be in the form
of: y = a(xb), where y is discharge in m3/s, a and b are coefficients,
and x is stage height in m.
e. Now you can calculate the discharge over the course of the season,
simply by plugging in the depth readings from the datalogger
measurements as the x variable in the equation. Create a discharge
column in the spreadsheet containing the datalogger depth
measurements, input the formula using the equation (substituting the
depth measurements for x). Then, plot the discharge value against the
date. This will give you a summer discharge profile.
F. TSS
1. Precombust the filters at 450°C for 4 hours in a muffle furnace to remove
any organic carbon on the filter surface.
2. Cool filters by storing in a desiccator overnight to prevent moisture buildup on the filter surface (Or place in an 60°C dryin g oven for about an
hour).
3. Measure the masses of the filters are measured on a 4 or 5-place balance
and record.
4. Store the filters in petri dishes with a label on which an identification
number (e.g. TSS09-001), and spaces for the recording of the station,
date, volume filtered is printed (See example below).
5. Record in the field book the identification number of the filter and the
volume of water that was filtered. Also record this information on the petri
dish label. Keep the filter in this dish until you get back to lab.
6. If you do not want to filter in the field, take a 2L sample of the water back
to the lab and us a vacuum set up to filter the water. Remember to record
the filter number and volume filtered.
7. Place the filter(s) into a numbered, pre-weighed aluminum weigh boat(s).
Record both the filter number and the weigh boat identification in a lab
book.
8. Place the weigh boat(s) into the 105°C drying ov en for at least twelve
hours. Cover the boat(s) with a sheet of aluminum foil to prevent foreign
material from falling onto the filter surface.
9. Place the boat(s) into a desiccator to cool. Record the weight of the room
temperature filter + weigh boat.
17 |
Arctic LTER Streams Protocol
10. Place the boat(s) into a preheated and stabilized, 550°C muffle furnace
for two hours.
11. Cool filters + weigh boat(s). Depending on the muffle furnace demand at
camp, either a) remove hot filter + weigh boats, and allow them to cool in a
GLASS desiccator, or b) turn off furnace and cool filter + weigh boat(s) in
furnace. Once filters are cool enough to handle, record the room
temperature weight of the filter + weigh boat.
12. Discard the filter.
13. Calculations
a. Using the six measurements below the total suspended sediments and
loss on ignition can be calculated.
• Tare weight of the dry, pre-combusted, numbered, unused filter,
(W filt, mg)
• The volume of water filtered (Vfilt, ml)
• Tare weight of the dry and empty weighing tin W tin, mg)
• Tare weight of the 105oC dried tin plus used filter (W dry, mg)
• Tare weight of the 550oC combusted tin plus used filter (Wcomb, mg)
• Tare weight of the combusted empty tin(W post, mg)
b. Calculations of TSS and LOI
• TSS = (W dry – W tin - W filt) / (Vfilt/1000) in mg TSS/L
• LOI = (W dry – Wcomb) / (Vfilt/1000) in mg LOI/L
• Weighing check = (W post – W tin) / (W tin)] * 100 or % change in tin
weight
G. Dripper
1. Phosphoric acid barrels should be ordered in April of each year because
they must be special ordered and take a while to get to Toolik. Once they
are shipped to Fairbanks ask the Logistics coordinator to hold them in
Fairbanks until June to prevent freezing. Unused barrels should be
shipped to Warm Storage for the winter.
2. Set up on the Kuparuk is done on 25 June every year. From 1985-1995
the dripper was at .56k, from 1996-2010 it was at 1.4k, from 2011 to
present there are two drippers: Big Drip at 0.0k, and Little Drip at 1.4k. A
helo flight should be scheduled before 25 June to sling the phosphoric
acid barrels to 0.0k in a fish tote, as well as the pump, batteries, and all
other materials needed. The dripper at 1.4k is walked from the parking
area at the bridge.
3. The dripper pump should be checked upon arrival to camp. The pump
should also be stored in Warm Storage for the winter.
a. check that all of the tubings fit and are long enough – use Teflon tape
to seal the screws to prevent air from getting in the line
b. check the timing – the dripper at 0.0k, “Big Drip” should drip at
2.4mL/min, and the dripper at 1.4k, “Little Drip” should be dripping at
1.2ml/min. – you can just use water to get it started
18 |
Arctic LTER Streams Protocol
c. Using the tubes and rope set up the dripper line so that the phosphoric
acid drips into the middle of the river. Use rebar and knots to get the
line taught and high over the river – the higher the better so that it is
less likely to get taken down in a high water event
d. check the dripper every day to make sure that there is plenty of acid
and it is dripping at the right rate. The acid barrel should be replaced
~1x/week
H. Macroinvertebrates
1. Surber samples
a. Using the standard Surber sampler collect two samples from each
macroinvertebrate collection station. Use a scrub brush to stir up the
sediment into the net of the Surber sampler. Rinse the sample out of
the net into a bucket. Pour the sample through a micron sieve. Pour
the sample into a whirl pack – large ones are necessary in the fertilized
reach because there will be a lot of moss in the samples.
2. Drift samples
a. record the number on the current meter, attach it to the drift net
b. set up the drift net perpendicular to the direction of flow
c. record the time
d. measure the height of the water on the net
e. after 15 minutes, take the net down, record the current meter number
f. rinse the sample into a bucket, and then through a sieve (same as with
the Surber sample), pour sample into a whirl pack
3. preservation
a. preserve all samples in the lab with ~10mL of formalin in the hood in
Lab 3.
I. Moss point transects
1. field equipment
a. 50m tape
b. view scope
c. field notebook and pencil
2. methods
a. Attach the free end of the field tape to one handle of the viewscope.
The other person (the ‘recorder’) plays the tape out at 20cm intervals
and simultaneously records the ‘hit’ identified by the caller.
b. Record the date, the start time (HH:MM), the station name, the interval
(20cm for all stations in the Kuparuk), the direction of movement
(TR>TL or TL>TR), and the relative location of the transect above the
immediate previous transect (e.g. +5m or +4 m).
c. Develop a simple alphanumeric key so that you can efficiently record
species without having to write down a lengthy name. It is most
efficient if two people work in the field as a team, with one person
moving across the stream calling out the observed cover types and
one person on the shore to record the observations.
19 |
Arctic LTER Streams Protocol
d. Record the ‘hit’ codes in the field book in columns. Start recording
down the page then jump up and to the right, going down a new
column until the transect is completed.
e. At each station, do a minimum of 5 (five) transects across the stream,
spaced evenly along the riffle. A spacing of 4-5 m works well for most
stations.
f. Note that if most stations are on the order of 20 m wide, a 20 cm
interval along transects will produce 100 points per transect and 500
points for a station with 5 transects. This is a desirable sampling
density.
• The point observations must be converted to percent cover (C%),
as follows:
C% = (Ni / Nt) * 100
where Ni is the number of observed points that match the class
type i (hits) and Nt is the total number of points observed. We use
an Excel spreadsheet to organize and automatically analyze the
data.
3. moss biomass
J. Fish
1. Permits
Permits must be applied for in the spring through AK Fish and Game. A report
must be filled out with specifications for what fish work will be done. A general
Toolik Area permit is always needed, other supplemental permits may also be
required (i.e. Anaktuvuk River Fire). If other parts of the project (i.e. Lakes) will
be doing any fish work be sure to have them fill out the information for their
section. IACUC forms are also needed each year. Some years it is just a
renewal, other years an entire permit must be written.
The AK Fish and Game permit requires that a state biologist must be contacted
(email is fine) before any fish work begins. Post-season reports must also be
filed, including a data report of all fish caught and a completion report with data
and analyses for all fish work. The data report is due 31 January of each year,
the completion report is due 1 June of each year.
2. Young of the Year (YOY)
a. Field equipment
• dip nets
• labeled YOY containers
• YOY hats
• polarized sunglasses
20 |
Arctic LTER Streams Protocol
• cooler with ice
• seine
• battery aerators
• backpack electrofisher
• salt
• orange insulated gloves
b. In the field
• At each station try to catch 10 YOY. YOY typically emerge earlier
on Oks than on the Kup by about a week. When they are small they
are usually found in slow moving backwaters and side channels
and are easier to catch. As they get bigger they move to faster
waters. The seine can be used to try to corral them into a smaller
area. It is easier to find them on sunny days, they often just don’t
come out from under rocks if it is cloudy. There are many
techniques that can be used – get creative and be patient. Do not
spend more than half an hour at any station.
• The electrofisher can be used when they are bigger, but you need
to add salt to the water to raise the conductivity enough for it to
work. Make sure everyone is wearing orange gloves and
waders that don’t leak before operating the electrofisher.
• When you catch them, place them into the containers with water,
then set the container in the stream (with a rock on top) to keep the
water cool. When you get back to the truck place the containers in
the cooler with ice. The battery powered aerators can be used to
keep the water oxygenated.
c. Return to the lab with live fish (check with Linda if that is the protocol
for the year, some years the YOY are kept for genetic information).
Most of the weighing and measuring equipment is in Lab 3.
• To weigh and measure the fish they should be anesthetized with
clove oil. At the beginning of the season make a dilution of clove oil
to be used for the summer.
o Put some water in a weigh boat, add one or two drops of clove
oil. Put one or two YOY into the weight boat until they are
anesthetized. Take one out at a time and measure with calipers.
Then place on the balance to weight. Record the length and
weight of each YOY.
o To help the YOY recover put them in a weigh boat with fresh
water and a drop or two of Stress Coat.
o Freshen the water in the clove oil and Stress Coat dishes as
needed
• Return all YOY to the appropriate river, including any casualties,
per the Alaska Fish and Game research permit.
3. Adult grayling
a. Field equipment
21 |
Arctic LTER Streams Protocol
•
spin reels and rods (check condition at the beginning of the season,
some may need replacing) or fly rods
• mepps spinner lures sizes #0 and #1 with barbs depressed and two
of three hooks cut off, or flies
• mesh holding bags
• mesh holding pens – 4x4x4 nylon-mesh with 4 rods of rebar per
pen to anchor pen in the stream
• Field balance (check batteries)
• pliers/Leatherman tool
• Weigh boat
• fish measuring board
• Square bucket
• Five gallon buckets (several)
• Sun shade
• Kit with tagging equipment
o syringes with rod plunger and tagging needles
o tags – check the protocol for the season – ½ duplex, full duplex,
Floy juvenile tags
o BioMark tag readers – reads both half and full duplex
check batteries and that it is working properly before going in the
field
o MS-222 for anesthetizing
o baking soda to raise the pH of MS-222 solution
o betadine solution – to sanitize the tagging needles between fish
b. What to do in the field – Fish!!
• there are several reliable areas – ask Linda or Bruce, they know
them well
• Set up the tagging equipment – keeping at least two 5 gallon
buckets – one for MS-222 and one for recovery, use the sun shade
to help protect the area from wind and as a staging area for
supplies
• Each person fishing should also have a 5 gallon bucket to hold and
carry any catches to the person tagging
• When each fish is caught anesthetize it
o tag it – for further tagging information see the PIT Tag Marking
Procedures Manual from the Columbia River Basin Fish and
Wildlife Authority
o measure total length of fish to the nearest 0.1cm from tip of nose
to bottom lobe of caudal fin (if caudal fin is missing or damaged
mention it in field notebook)
o measure wet weight to nearest gram using the portable field
balance
o record tag number, length, weight, and location in the field book,
as well as any other appropriate comments
22 |
Arctic LTER Streams Protocol
•
•
•
o Place the fish in the stream. Hold the fish by the caudalpeduncle, head into the current, until the fish has had an
opportunity to recover from the anesthesia. Release the fish.
if there are several fish at once keep some in the mesh holding
bags while others are being worked up
if fish are to be weighed on an empty stomach, keep them in the
holding pen overnight – return the next day to tag and weigh
if stomach contents are being analyzed
o While it is still anesthetized, hold the fish with its mouth over the
catchment pan.
o Fill a 60-cc syringe (loaded with a 13-gauge needle, tipped with
rubber tubing to prevent scratching) with water and carefully
insert tubing down the fish’s esophagus.
o Inject the water, forcing the fish to egest its stomach contents
through the mouth and into the pan. Gentle pressure on the
fish’s stomach helps to induce egestion.
o Filter stomach contents through the 100-µm nylon mesh.
o Transfer the stomach contents from the mesh into a 250-ml
plastic bottle (use the funnel to make this easier). Label and
preserve stomach contents in 95% ethanol.
o Place the fish in the stream. Hold the fish by the caudalpeduncle, head into the current, until the fish has had an
opportunity to recover from the anesthesia. Release the fish.
o Return stomach contents to camp. Store in a sturdy box and
ship or hand-carry to RESL for analysis.
c. weir
• set up
II.
Lab Methods at Toolik
The “nutrient” RA helps in Wet Lab to make reagents and standards for
everyone. These procedures include the methods used by the Nutrient RA as
well as the procedures followed to analyze samples.
A.
Soluble Reactive Phosphorus
The method used for soluble reactive phosphorus determination is based on
Parsons et al. 1984. It involves the reaction of phosphorus with molybdate,
ascorbic acid, and trivalent antimony. The molybdic acids are reduced to a bluecolored complex which is then read for absorbance on a UV spectrophotometer.
A 1 cm cell is used in the spectrophotometer.
23 |
Arctic LTER Streams Protocol
1. Mixed Reagent for Phosphate Analysis:
This MUST be made the day you want to use it – it is only stable for 6 hrs. Solns
a, b, and d can be prepared ahead of time in bulk. Soln c must be made daily.
Use pre-labeled bottles, volumetrics and graduated cylinders.
ADD TO CLEAN CALIBRATED (500 uL) RE-PIPETTER in the following order:
a. 50 mL Ammonium para Molybdate Solution
b. 125 mL Sulfuric Acid Solution
c. 50 mL Ascorbic Acid Solution *MUST BE FRESH*
d. 25 mL Potassium Antimony-Tartrate Solution
This will be enough for 500 samples.
2. Stock Solutions:
There are a set of bottles/graduated cylinders that are reused to make these
solutions. Keep them labeled and don’t use for other purposes.
a. Ammonium para Molybdate Solution:
• Dissolve 15 g Ammonium para Molybdate ((NH4)6Mo7O24.4.H2O) in
500 mL DI water. Store in pre-labeled amber bottle – stable
indefinitely.
b. Sulfuric Acid Solution:
• **Safety glasses, gloves, lab coat and closed toed shoes required.
Do not combine ingredients into a volumetric flask as the reaction is
exothermic and will stretch the volumetric**
•
Add 70 mL of concentrated H2SO4 (Sulfuric Acid – 4.8 N) to 450 mL
DI. Allow solution to cool. Bring to 1 L. Store in glass amber bottle.
c. Ascorbic Acid Solution – MAKE DAILY
• Dissolve 2.7 g Ascorbic Acid in 50 mL DI (can use graduated
cylinder)
d. Potassium Antimonyl – Tartrate Solution:
• Dissolve 0.34 g Potassium Antimony in 250 mL DI water (warm if
necessary). This solution is stable for many months.
3. Standards
24 |
Arctic LTER Streams Protocol
a. Stock A Solution-1000mM PO4-P
•
Dissolve 0.136 g of anhydrous potassium dihydrogen phosphate
(KH2PO4, m.w. = 136.07) in approximately 900ml of deionized
water contained in a 1 L volumetric flask (note all salts should be
dried in oven prior to weighing).
•
Dilute the solution to the mark with deionized water and mix it
well.
•
Transfer this solution to an amber bottle.
•
Add 1ml chloroform for preservation. This solution is stable for
many months. Refrigerate when it is not in use.
b. Stock B Solution-100mM PO4-P
•
Prepare this solution weekly.
•
Using a volumetric pipet or a calibrated automatic pipet, add
10.0ml of Stock A to approximately 90 mL of deionized water
contained in a 100ml volumetric flask.
•
Dilute the solution to the mark with deionized water and mix it
well.
•
Add 1ml chloroform for preservation and store in an amber
bottle.
c. Working Standards
•
Prepare these solutions daily.
•
Use adjustable, micro liter pipettes to add the designated volumes
listed in the following table.
•
Calibrate the pipet for each required volume. (NOTE: The
standards for each day are presented in bold typeface. The other
concentrations are included for reference, if needed.)
•
Prepare each standard by adding the required amount of stock to
the required volume volumetric flask containing deionized water.
•
Standards for both Ammonium and Phosphate can be made in
the same volumetrics.
•
After adding standards for both methods, dilute each to the mark
with deionized water and mix well. Keep these solutions tightly
sealed.
4. Sample Prep Procedure
25 |
Arctic LTER Streams Protocol
a. Sign up for a date and time on the door to the PO4 room specifying the
number of samples and volume of reagent needed.
b. Make sure the sample tubes are clean and dry
c. Label the rack with tape and station names for ease when reading
samples.
d. When ready to run samples, calibrate the pipette for 5mL
e. Rinse the pipette tip 3X’s with DI
f. Rinse the tip 1X with sample
g. Pipette 5mL of sample into clean tube
• run triplicates of each sample
• Also run triplicates of check standards for every 10 samples
o the Arctic LTER Streams group uses the Nutrient RA standard
curve, but a standard curve can also be run if needed
• Rinse 3X’s with DI and 1X with next sample before each sample
h. Dispense reagent from repippeter a few times to make sure there is no
air
i. Add 0.5mL of working reagent from the repipettor to each sample
j. Vortex each sample
k. cover with saran wrap
l. let sit for ~30-90 min in the light
5. Spectrophotometer reading (follow the Nutrient RA’s instructions)
a. Make sure the waste tube is placed in correct waste container
b. Flush with degassed DI a few times before running samples
c. Read DI to get a stable number
d. Once DI is stable begin reading samples
e. Vortex samples before reading
f. Read each sample, recording the numbers in the notebook
g. If there are errors, check with the Nutrient RA about what should be
done
6. Vial Cleaning
a. Pour samples into phosphorus waste container
b. Rinse tubes once with DI into the waste
c. Rinse tubes once with DI into the sink
d. (Remove tape from rack) put rack in acid bath
e. Rinse with DI 3X’s
f. Dry upside down
26 |
Arctic LTER Streams Protocol
B.
Ammonium using the OPA method
1. Overview
The OPA method began being used in 1999. A year of comparison between
the phenol method and OPA method occurred that year. After 1999, only the
OPA method has been used.
Fluorescence is produced by the reaction of OPA with ammonium.
Fluorometry is sensitive and simple so it seems to be a good way to measure
ammonium, particularly at low levels. Details of methods, reagents, etc are
given in Holmes et al. 1999 (CJFAS 56:1801-1808). This document
supplements the manuscript and is intended to give a quick, user-friendly,
informal overview of the procedure. It also details a variation of the method
not discussed in the manuscript, which uses 2 mL sample and 8 mL working
reagent.
a. Background Fluorescence
All samples auto-fluoresce to some degree. This BF must be subtracted from
the observed sample fluorescence in order to quantify ammonium
concentration. If it is found that BF doesn’t change through the water column
or down a stream transect it may be possible to take fewer BF
measurements. If it does change however you will need to take a BF each
time a sample is taken. In surface waters around Toolik Lake, ammonium
concentrations tend to be very low and background fluorescence is relatively
significant. Therefore, it is important to accurately quantify BF. In our limited
experience so far, BF is relatively constant in a given water-body on a given
day (for example, Toolik Main station or Kuparuk River transect), but BF
varies across stations (and maybe temporally). Therefore, BF does not need
to be sampled at every station within a given “station”, but must be sampled at
each stream. Another example: On June 23, 1999, BF was essentially
constant at 11 Kuparuk River stations, but differed significantly in Hershey
Creek, a small tributary to the Kuparuk River. If BF had not been measured
in Hershey Creek and instead the Kuparuk BF was used, the Hershey Creek
ammonium result would have been erroneous.
b. Matrix Effects
A summary of matrix effects are included below, but in 04 the lakes groups
decided that the ME of Toolik Area lakes were close enough to DI that we
could simply run a nutrient standard curve in DI and no longer utilize standard
additions. The difficulty and inherent error of spiking with such small volumes
of standard by unseasoned SRA’s also led to this decision.
27 |
Arctic LTER Streams Protocol
OPA and ammonium react differently in different waters. In DI water, a given
amount of ammonium tends to produce more fluorescence than it would in
lake or river or soil solution samples. To quantify ME and correct for it,
standard additions are done to samples and compared to DI water standards.
For surface waters around Toolik Lake, we have been spiking samples with
50 ul of 50 uM ammonium stock solution to quantify ME. In general, ME have
been around 5-25 %. This correction is generally on the order of 0.01-0.03
uM for surface waters around Toolik Lake, but will be greater when
ammonium concentrations are greater. Therefore it is important to note that
for higher ammonium values a larger spike is required to assess ME. As with
BF, ME appears to be relatively constant within a given water-body but will
probably vary across stations and maybe temporally.
2. Reagents and Equipment
The reagents are made by the nutrient RA. Sign up for a time to react your
samples and a time to read your samples. The OPA room should remain dark
at all times. OPA is light sensitive.
The following is enough for about 48 liters of WR and 4 liters of Borate Buffer.
Reagent
Size
Sigma cat #
Sodium sulfite
250 g S 4672
Sodium borate
2.5 kg S 9640
Orthopthadialdehyde 100 g P 1378
ETOH
2.5 l
3. Preparation of Stock reagents
a. BORATE BUFFER (BB): Borate buffer without the sodium sulfite or the
OPA is used to evaluate the sample background fluorescence (BF).
b. SODIUM SULFITE: Next prepare the sodium sulfite solution (2 g
sodium sulfite to 250 mL DI water)
c. ORTHOPTHADIALDEHYDE (OPA): add 8 g OPA to 200 mL ethanol
(keep this solution as dark as possible); shake vigorously until OPA
dissolves.
d. WORKING REAGENT (WR): Working reagent appears to be stable
for months, and its blank fluorescence decreases over time, so it is
best to make WR in large batches and let it age. We make WR
batches of about 4 L in 1 gallon brown Nalgene® bottles (these bottles
actually hold about 4.4 L).
•
28 |
To a clean 4-liter bottle (pre-react, or just rinse with DI if previously
used for WR), add approximately 3 L DI. Then add 160 g sodium
borate, cap, and shake vigorously until your arms are tired, then
Arctic LTER Streams Protocol
rest, then do it some more, and add 20 ml of sodium sulfite solution
to the 1 gallon jug with DI and sodium borate already added.
Shake the jug some more. Finally, add 200 ml of OPA solution to
the 4-liter jug. Shake some more, and then add DI until the bottle is
nearly full – about 1 inch from the top. Shake a bit more, let age for
at least a few days if possible, and then the WR is ready to use.
4. Fluorometer and filters
Optical Kit for Holmes NH4 Method
See web http://www.turnerdesigns.com/t2/doc/appnotes/s_0025.html
The Ammonium Optical Kit (P/N 10-303) includes two Near UV Mercury
Vapor Lamps, 350nm excitation filter (310-390nm), a 410-600nm emission
filter, a 1:75 Attenuator plate, and a 10-300 Reference Filter (>300nm). The
10-303 Optical Kit works in the 10-AU Digital Field Fluorometer, TD-700
Laboratory Fluorometer and the Model 10 Analog Fluorometer and also
includes a reference filter (P/N 10-300).
5. Calculations
Four fluorescence values and the slope of a standard curve are needed to
calculate uM NH4
a. 1:4 DI H20/Working Reagent, DI WR
b. 1:4 DI H20/Borate Buffer, (for to correct for background fluorescence)
DI BF
c. 1:4 Sample /Working Reagent, Samp WR
d. 1:4 Sample/Borate Buffer, (for to correct for background fluorescence)
Samp BF
e. Slope of 0-3uM NH4 Std Curve
Calculating uM NH4
uM NH4= {Samp WR-(DI WR+(Samp BF-DI BF))}/{Slope of curve}
6. Procedure
a. Pre-reaction of vials (only need to do at beginning of season or with
new vials):
• New vials: Rinse 3x with DI, fill with 8 ml dilute working reagent and
let sit for 3-24 hours.
• Old vials: Rinse 1x with DI, fill with 8 ml dilute working reagent and
let sit for at least 3 hours.
b. Dump WR into NH4 waste, rinse with 2-3 ml dilute (if available) or new
working reagent
c. Fill with 8ml new working reagent.
d. Before setting up a run:
• Dump old samples from vial into NH4 waste.
• Rinse vials with DI, dump into NH4 waste
29 |
Arctic LTER Streams Protocol
•
•
•
Rinse vials with 2mL borate buffer, dump into NH4 waste
Fill WR vials with 8 ml good working reagent. Use the same batch
of working reagent for all samples and standards in your run.
Fill buffer vials with 8 ml new borate buffer (don’t need to rinse).
e. After you return from the field with your samples:
• Let standards and samples warm to room temp if not already.
• Pipette 2 ml of each standard (made by nutrient RA) into their
respective vials (each standard and sample has one buffer vial and
2 working reagent vials).
• Change pipette tip after pipetting standards and before pipetting
samples. You can use the same pipette tip for all samples, but rinse
the tip 1x with DI and 1x with the next sample first.
• Pipette 2ml of samples into each row of numbered vials. Record
which sample is in which row of vials.
• Do all pipetting in the dark. OPA working reagent is sensitive
to light.
• Shake samples and store in the dark for 5-24 hours. Record the
time the samples were shot up.
f. Reading samples• Verify that fluorometer is on – if not push the red button. Let it warm
up for about 30 minutes before reading samples
• Use two tubes – one for buffer and one for working reagent
• Rinse 13x100mm borosilicate tube with DI
• Rinse 13x100mm borosilicate tube with a small amount of sample
from scint vial – vortex
• Pour sample from vial into borosilicate tube - vortex
o Rinse exterior of glass tube with DI and wipe with kimwipe/tissue
to remove any fingerprints or residual WR.
• Place tube into fluorometer, replace black cap
• Allow reading to stabilize (about 5-10 seconds), press [*] and read
average fluorescence value.
• reuse borosilicate tubes, rinsing 1x with DI and 1x with sample in
between.
• Use new tubes if going from a high standard of sample to low.
• Use separate tubes for buffer samples and WR samples.
C.
Chlorophyll a
1. Reading Samples on Turner Designs10-AU Fluorometer
TD-10AU should already be calibrated and ready for use, for more
information regarding calibration, lamp/filter arrangements, etc. refer to users
manual or Fluorometer use protocols.
30 |
Arctic LTER Streams Protocol
*Safety Precautions, chronic exposure to acetone can cause health problems
(see MSDS). Always wear gloves, eye protection, and make sure snorkel
hood is working properly.
a. Verify TD fluorometer is on, if not press red button at front of unit and
allow to warm up for 20 minutes.
b. Use the Chlorophyll notebook to record data
c. Place chlorophyll solid standard in fluorometer, replace black cap, wait
2 minutes for reading to stabilize, press [*] to read average and record
values for both the high and low std. To switch between the two simply
turn 180 degrees till solid slides into place. If readings deviate from
mean std. readings by more than 10%, the unit may need recalibration
(see Lakes or Nutrient RA).
d. Read and record resting fluorescence with no tube in fluorometer. It
should be around 0.328. This will help you catch problems associated
with changes in lamp intensity. A second check is a clean glass tube
which has a fluorescence around 3.32.
e. Read and record fluorescence of two acetone blanks (90% acetone
w/MgCO3) in 13x100 glass tubes
f. Reading samples - Carefully transfer extract from Falcon tube to a
clean 13x100mm borosilicate tube (leave last 0.5ml of acetone, filter,
and any particles in the Vulcan tube).
g. Wipe exterior of glass tube with kimwipe/tissue to remove any
fingerprints or residual acetone.
h. Place tube into fluorometer, replace black cap
i. Allow reading to stabilize (about 5-10 seconds), press [*] and read
average fluorescence value, record preacidification value (Rb).
j. Using calibrated pipette, transfer 300ul 0.1N HCL (or 100ul 0.3N HCL)
into the sample you just read, mix by inversion (place Saran wrap over
end, hold with thumb) or by using disposable glass pipette to “mix”
k. Let stand for 2 minutes (its easiest to work out a system where you
read a row unacidified, then acidify and let stand while reading 2nd row,
acidify, etc, etc)
l. Repeat steps 7-10, record post acidification fluorescence (Ra)
m. After recording both readings, dump contents of glass tube into
acetone waste container, rinse with small amount of DI and acetone,
let dry.
n. When finished with samples rerun and record low and high chl. solid
standard values (let readings stabilize about 2 minutes)
o. Make sure all glass tubes are rinsed and disposed of properly, place
cap on acetone waste bottle, clean up kimwipes, pipettes, acid, etc.
Leave fluorometer on for next user
p. Remove filters from Vulcan tubes dispose into filter bin, rinse tubes
with DI, and acetone.
q. See equations below to determine chlorophyll a and pheophytin a
concentration.
31 |
Arctic LTER Streams Protocol
Chlorophyll a
C E, C = Chlorophyll a concentration in the extract (usually 10ml)
C E, C = Fs(r/r-1)(Rb-Ra)
Where:
C E, C = corrected chlorophyll a concentration (ug/L) in the extract solution
analyzed
Fs = response factor for the sensitivity setting S
r = the before to after acidification ration of a pure chl a solution (Rb/Ra)
Rb = fluorescence of sample extract before acidification
Ra = fluorescence of sample extract after acidification
C S, C = corrected chl a concentration (ug/L) in the whole water sample
C S, C ={(C E, C x extract volume in L x Dilution Factor)/(sample volume
in L)}
Pheophytin a
PE = pheophytin a concentration (ug/L) in the sample extract
PE = Fs (r/r-1) (rRa - Rb)
PS = pheophytin a concentration (ug/L) in the whole water sample
PS = (PE x extract volume in L x dilution factor)/(sample volume in L)
Note: if no dilution took place, then DF equals 1
32 |
Arctic LTER Streams Protocol
III.
Lab Methods at the Rubenstein Ecosystem Science
Laboratory (RESL)
A. Nitrate
QuikChem Method 31-107-04-1E for use on the Lachat (previously, QuikChem
Method 31-107-04-1C was used)
This is a condensed version of the QuikChem method with details specific to
working at the RESL on Arctic samples. It is advised to refer to the Lachat
method for further information about this analysis. Samples are passed through a
copperized cadmium column to reduce nitrate to nitrite. The nitrite (both original
and reduced nitrate) is determined by diazotizing with sulfanilamide and coupling
with N-(1-napthyl)-ethylenediamine dihydrochoride to form a highly colored azo
dye which is measured colorimetrically.
1. Keep up to date notes after every use of the Lachat. This will help you to
keep track of problems, and ways you have solved them in the past.
Reagents – prep the day before getting started on the Lachat
a. 15N sodium hydroxide
b. 150 g NaOH into 250 mL water – the solution will get HOT – caution!
Make it in the fume hood!
c. Ammonium chloride buffer – pH 8.5
• dissolve 85.0 g ammonium chloride and 1.0 g disodium
ethylenediamine tetraacetic acid dihydrate in a large beaker in ~800
mL water.
• calibrate the pH meter with standards
• adjust the pH of the ammonium chloride buffer solution by slowly
adding sodium hydroxide while the pH probe is in the solution.
• once everything is dissolved, pour into a 1L volumetric flask – rinse
the beaker with DI into the flask to make sure all particles are in
solution. Fill to mark with DI.
d. Sulfanilamide color reagent
• to a 1L volumetric flask add ~600mL water.
• Sulfanilamide is light sensitive – wrap flask in aluminum foil
• add 100mL 85% phosphoric acid, 40.0g sulfanilamide and 1.0g N(1-naphthyl) ethylenediamine dihydrochloride (NED). Stir to
dissolve for 30 min. Dilute to the mark and invert to mix. Store in a
dark bottle – it is stable for ~1 month.
e. Nitrate and nitrite standards
• To make primary standards (71.4uM N or 1000mg N/L)–
o The primary standards for both NaNO3 and KNO2 need to
be placed into an oven at 100C for two hours to burn off any
moisture weight. All standards are made from the primary
standard.
33 |
Arctic LTER Streams Protocol
o Nitrate: Dissolve 6.04g NaNO3 into 1 Liter of deionized
water.
o Nitrite: Dissolve 6.048g KNO2 into 1 Liter of deionized water
STD Volume of stock (71.4uM
Label diluted into 100mL
A
100ml of Stock in 250ml
Volumetric Flask.
B
50 ml of Standard A in
100ml Volumetric Flask
C
50ml of Stock
D
2.5ml of Stock
E
0.5ml of Stock
F
0ml of Stock (100ml DI)
Concentration
28.6uM
14.3uM
7.15uM
1.788uM
0.36uM
0uM
•
Determine cadmium column efficiency (efficiency is determined by
comparing the nitrate to nitrite ratio. It was determined that a
beginning column efficiency should not be less than 90%. Don’t use
the numbers after the column efficiency is below 85%).
2. Getting ready to run samples
a. get samples out to thaw – a water bath can be used, or spacing them
out on the bench
b. Run DI through the tubes for ~10 min before running
c. Load the method to be used – check the timings, concentrations, and
DQM
• the DQM is the QAQC for the program – it is good to run check
standards once ~10-15 samples, including a column efficiency
check.
• Run replicates 1/10-15 samples as well to check the machine
d. Load the table to be run – fill in the sample names and order
e. Begin running reagents through the tubes
f. Put the column on – be careful not to get air in it! Columns contain
cadmium, which is toxic! Cadmium columns began being purchased
(50237A from the Hach website) instead of recopperizing and packing
because they can get ~1,000 samples, which saves time and contact
with cadmium. The columns are stored in buffer – this seems to help
keep air out and make them last longer.
g. Run a standard curve to check baseline, standards, and column
efficiency. If everything looks ok, samples can be run. If the samples
are not ready yet, turn the column off and set the pump to minimum
speed to reduce waste of reagents.
h. To continue prepping the samples, pour samples into clean
borosilicate tubes. We pour samples because the centrifuge tubes
often break. 60mL of sample is available, so that if samples need to be
rerun there is plenty of sample, however, this means that it can take a
34 |
Arctic LTER Streams Protocol
while for the entire sample to thaw. Be patient, it is best if they are run
at room temperature.
i. Before running the tray, make sure to clear the standard curve. Once
everything is ready, hit Run tray (be sure to turn the column back on
and set the pump to normal speed). Stay with the machine for a little
while to make sure everything is running smoothly. Check to make
sure the arm is working properly, check to make sure the tray is lined
up correctly so the needle does not get out of line and cause air to be
sucked.
j. If air does get sucked turn the column off immediately. If air gets in the
column you can try to blow it through by pressing the max speed on
the pump. If air gets in, stop the run, and let buffer run through for a
little while – this sometimes help to work it out. Check the efficiency
again, and try the run again.
3. If samples have very low concentrations of nitrate there are other methods
available that have a lower detection limit, the one used has been
historical.
4. If samples have negative concentrations spike additions can be run to
determine if there is a matrix effect. If recovery is high (<95%) then it is
likely that there is simply no nitrate in the sample.
a. Spike additions to determine if there is sample interference
• The volume of the spike additions also varied depending on the
sample volume in the tube. Two equations are necessary:
o (Sample volume) / (Stock concentration/ Desired Concentration)
= Spike Volume
o (Stock Concentration * (Spike Volume /(Sample Volume + Spike
Volume))=True concentration
• The first equation gives an approximate spike volume to add to the
sample. The second equation gives the true concentration of the
spike based on the approximate volume. The desired
concentrations for this procedure are 0.50 µM for the low spike, and
2.0 µM for the high spike. The true concentrations were 0.49 µM
and 1.92 µM, so for the 8 ml : 8ml example above, the low spike
was 80 µL and the high was 320 µL.
o In 2008 and 2009 spike additions were done (both samples and
the standard curve were spiked)
A high spike had 0.5mL of 50uM standard added to 5.0mL
sample to produce a concentration of 4.54uM
A low spike had 0.125mL of 50uM standard added to 5.0mL
sample to produce a concentration of 1.22uM
35 |
Arctic LTER Streams Protocol
B. TDP
The determination of total dissolved phosphorus is done by liberating organic
phosphorus as inorganic phosphate through oxidation by persulfate. The total
phosphate can then be determined using the molybdate method.
Stock Solutions – prepare ahead of time
Solution
Concentration
Ammonium
25g/250 mL DI
heptamolybdate
tetrahydrate
Potassium antimonly
tartrate
Sulfuric acid
Acidified DI for working
standards
Primary phosphorus
stock – potassium
phosphate anhydrous
Secondary phosphorus
stock
Primary Sodium
pyrophosphate
decahydrate digestion
check standard
Secondary sodium
pyrophosphate digestion
standard
Ascorbic Acid solution
for Working Reagent 1
Stability
Stable for months in dark
bottle, check for
precipitate and stir before
use
Stable for months in dark
bottle
5g/200 mL DI
9 N H2SO4 = 250 mL 36 N
H2SO4 made into 1L
0.012N = 12mL 1N HCl
made into 1L
10,000 M = 1.36g
KH2PO4 to 1L DI
Use to make working
standards
Stable for months
refrigerated
50 M P = 5mL primary
Stable for weeks
stock diluted to make 1L
refrigerated
1.33 g/ 1L = 10,000 M P/L Stable for weeks
refrigerated
50 M P = 5 mL primary
stock diluted to make 1 L
Stable for weeks
refrigerated
2g ascorbic acid dissolved
into 10 mL DI
Stable for the day:
(ensure it is freshly made
daily)
Digestion Solution - prepare fresh daily
Solution
Persulfate solution
36 |
Contents
Potassium persulfate
K2S208
Arctic LTER Streams Protocol
Ratios
2.5 g/ 50 mL DI
Difficult to dissolve –
use a little heat if
necessary – DO NOT
overheat
Standard dilutions – make fresh daily in dedicated 100 mL volumetric flasks,
digestion check standards can be made in a similar way
Concentration
Dilution from Secondary (2o) 50 M
Phosphate Standard with acidified
DI
0.00 M P
0.00 mL 2o + 100 mL acidified DI
0.05 M P
0.1 mL 2o + 99.9 mL acidified DI
0.10 M P
0.2 mL 2o + 99.8 mL acidified DI
0.25 M P
0.5 mL 2o + 99.5 mL acidified DI
0.50 M P
1.0 mL 2o + 99.0 mL acidified DI
1.0 M P
2.0 mL 2o + 98.0 mL acidified DI
1.5 M P
3.0 mL 2o + 97.0 mL acidified DI
2.5 M P
5.0 mL 2o + 95.0 mL acidified DI
Concentration calculation M1V1 = M2V2
(50
M P 2o stock solution)*(V1) = (x Molarity)*(100 mL)
V1 =
(x Molarity)*(100 mL)
50 M P 2o stock solution
Procedure
1. Prepare for Analysis:
a. Create spreadsheet with sample info and label autoclave and regular
racks
b. Organize 100ml Volumetric Flasks to contain each concentration of the
curve as prescribed in the “Standard Dilution” table above. Create
dilution with check standards in the same fashion (typically of 0.1 M
and 0.25 M).
c. Put each of the standards in a prescribed beaker or sealable jar for easy
pipetting
d. Ensure test tubes for standards are labeled as the first seven on the rack.
Include two digestion check standards in two additional test tubes after
the standards.
2. Pipette 7.5 mL of standards, digestion check standards, and sample into
Kimax screw top tubes
a. Include a 0.0 uM P and mid-range check standard at the end of each
sample rack row
b. Include at least one lab replicate in each sample rack row
c. Follow general pipetting rules – rinse tip with DI three times, rinse once
with sample, pipet sample into tube
3. Add 0.188 mL potassium persulfate solution to each tube, vortex
4. Cap tubes tightly
5. Check levels of liquid in tubes – etch level on some
6. Fill bottom of autoclave with water
37 |
Arctic LTER Streams Protocol
7. Check other liquid levels in autoclave
8. Adjust settings on autoclave – 90 min at 105oC
a. Place racks in tray
b. Autoclave for 90 min
9. Remove tubes from autoclave and let cool
a. Refrigerate if not running until the next day
Working Solutions – prepare fresh daily – lasts about six hours
**** Be sure to use chemicals in correct concentrations as labeled on first
page ****
Solution
Contents
Ratios
Working Reagent 1
9 N Sulfuric acid
1:1
Adjust volumes as
solution and ascorbic
10 mL 9 N sulfuric acid
needed
acid solution
+ 10 mL fresh ascorbic
COVER IN FOIL!!
acid solution (2g/10ml)
Working Reagent 2
9 N sulfuric acid
70:25:4
Adjust volumes as
solution: Ammonium
14 mL 9 N sulfuric acid
needed
molybdate stock:
+ 5 mL ammonium
potassium antimony
molybdate solution + 0.8
tartrate stock
mL potassium antimonl
tartrate solution
10. Run standard curve
a. Add .15ml WR1 – vortex
b. Add .15ml WR2 – vortex
c. Stand 30 min (covered with foil or saran wrap, but in the light)
d. Run on spec at 885nm
11. When standard curve is acceptable run samples
a. Add .15ml WR1 – vortex
b. Add .15ml WR2 – vortex
c. Stand 30 min
12. While samples are reacting
a. Make non-digested calibration curve (don’t need organic standards or to
acidify)
i) Pipette 7.5ml potassium phosphate standard solution into clean vials
ii) Add .15ml WR1 – vortex
iii) Add .15ml WR2 – vortex
iv) React for 30 min
13. Run samples on Spec
14. Run non-digested curve on spec.
15. Clean all volumetric flasks, beakers, and sealable jars. If reusing volumetric
flasks or sealable jars for another day, clean and fill with DI water. Let DI
water sit until glassware is used again.
38 |
Arctic LTER Streams Protocol
Chemicals list
Potassium persulfate (K2S2O8) – J. T. Baker 3239-01
Potassium Phosphate (KH2PO4) – Fisher P285-500
Hydrochloric Acid – Trace Metal Grade – Fisher
Sulfuric Acid – Trace Metal Grade – Fisher
L-Ascorbic Acid – Fisher A61-100
Potassium Antimony Tartrate C8H4K2O12Sb2*3H20 – J.T. Baker
0864-4
Ammonium Molybdate (NH4)6Mo7O24*4H20 – Fisher A674-500
Supplies
39 |
Volumetric flasks for standards
Sealable jars for standards
Pipettes
Pre-made solutions to make working reagents
Clean tubes, racks, caps
Arctic LTER Streams Protocol
C. TDN/DOC
1. Beginning in 2005 the Shimadzu TOC-L was used for TDN and DOC.
Previously; TDN was determined by persulfate digestion and analyzed on
the Lachat.
2. The TOC-L uses carrier gas, which is used to supply oxygen at
150mL/min to the combustion tube filled with oxidation catalyst. This tube
is heated to 680 degrees C and samples are burned in the combustion
tube to form carbon dioxide. The carrier gas, containing the carbon dioxide
created from the combustion, flows into a dehumidifier, where it is cooled
and dehydrated. The gas then goes through a halogen scrubber before it
reaches the cell of a non-dispersive infrared NDIR gas analyzer. This
analyzer detects the carbon dioxide, and the analog detection signal of the
NDIR forms a peak. The area of this peak is measured by a data
processor, and the peak’s area is proportional to the concentration of the
TC in the sample. To detect TN, the sample also goes through a
combustion tube at 720 degrees C. The sample decomposes at this
temperature to become nitrogen monoxide. The carrier gas carries the
nitrogen monoxide to a dehumidifier, where it is dehydrated. Then it enters
a chemiluminescence gas analyzer, where the nitrogen monoxide is
detected. The detection signal from the analyzer genterates a peak which
are proportional to the concentration of the TN in the sample.
3. Running the Shimadzu TOC requires a combustion column, O2 tank, and
standard.
a. The combustion column may need to be repacked with reconditioned
pellets. The supplies should be in the drawer next to the machine. A
sample column is available to know how much of each layer is
required. Quartz and glass cotton are used at each end of the column.
PROCEDURE
1. Prepare Needed Stock Standards.
a. Primary Stock Solution: 160,000uM C, 20,000 uM N.
i. To prepare, add 4.08g KHP and 2.02g KNO3 to DI water.
Dilute to mark with DI water.
b. Secondary Working Solution.
i. Add 10ml of Primary Stock Solution to DI water. Dilute to
mark with DI water.
2. Prepare TOC for Analysis.
a. Open Air Gas, take note of gas pressure. Ensure it is at least above
600psi to prevent running out of gas mid-run. Check the pressure
on the tank and the flowmeter in the machine to make sure flow is
appropriate.
b. Add Secondary Working Solution to the small loose tubing coming
out of the left hand side of the TOC. This sampling tube is known
40 |
Arctic LTER Streams Protocol
3.
4.
5.
6.
7.
8.
41 |
as “Vial 0” to the computer program. There should be another
piece of tubing going to the ASI auto sampler.
c. Check to make sure adequate levels of acid and DI water are
present within the analyzer. Fill as needed. Check the manual to
see what should be checked daily, and occasionally. Ensure that
everything is ready for analysis.
d. Turn on TOC and ASI auto sampler, allow it to warm up.
Open the “TOC-L Sample Table Editor”.
Create a New Sample Table.
Insert a “Conditioning Sample”. This will help “warm up” the TOC to get
ready for analysis.
a. Right click on the first cell on the Sample Table.
b. Click Insert-Sample
c. Choose “20120216_CN_ebs.met” as the Method.
d. Continue by clicking “Next”, through the current settings. Current
Settings should be set as: Manual Dilution:1; No. of
Determinations:1; Units:mg/L; Injection Volume:150; Expected
Conc. Range: 400; SD Max:0.1000; CV Max:2.00%; No. of
Washes: 2; Auto Dilution:1; Sparge Gas Flow:90; Sparge Time:
01:30; Acid Addition: 1.5%. Click Finish when at end.
e. Right Click on Sample Row Number. Go to Measurement Settings.
i. Click on the NPOC tab, Change the Number of Dilutions
from 3/5 to 15/15.
ii. Click OK.
Insert the TOC calibration curve for the run.
a. Right click on the second cell on the Sample Table.
b. Click Insert- Calibration Curve.
c. Choose “NPOC_19mgL.cal”. Click Open.
Insert the TDN calibration curve for the run.
a. Right click on the third cell on the Sample Table.
b. Click Insert- Calibration Curve.
c. Choose “TN_2.8mgL.cal”. Click Open.
Insert Samples
a. Right click on the forth cell on the Sample Table.
b. Click Insert-Multiple Samples
c. Choose Choose “20120216_CN_ebs.met” as the Method. Click
Next.
d. Choose Number of Samples to insert. Keep in mind that you will
also need check standards and DI samples throughout the run.
Click finish.
Arctic LTER Streams Protocol
9. The Vial Settings tab will open up. Change all vial numbers for the
conditioning sample, and standard curves to “0”. Do this by clicking on the
cell and hit enter after each entry. After completion, click OK.
10. Enter sample numbers into the spreadsheet, in the place which they will
be within the ASI-L auto sampler. It is suggested that after each 10
samples, a check standard and DI water test should be analyzed. Ensure
that all samples are poured into the vial and then it is sealed with the lid
and septum. DI should be within the ASI-L auto sampler with the samples.
The check standard should be set as “Vial 0”, and should be set to an
auto-dilution of 8 times. As you enter samples you will have to change the
Vial numbers in which they will be. You will have to go to the vial settings
to adjust these numbers. The vial settings can be accessed by clicking the
“birthday cake” like icon, which is supposed to look like the ASI-L auto
sampler, in the upper right hand side of the Sample Table Sheet.
11. Connect to the TOC-L analyzer within the program. Click on Monitor to
ensure that the TOC has warmed up to the appropriate temperature and
the proper amount of air pressure going into the system. If all parameters
are met, click on “Start”. Choose if you want the analyzer to shut down or
stay on after analysis, and click “Start”.
12. Allow the Analyzer to complete. This may take a few hours or overnight
depending on how many samples are being analyzed.
13. Acid Wash all TOC vials, and then ash the vials at 500 degrees Celsius for
2 hours.
42 |
Arctic LTER Streams Protocol
D. Particulate Phosphorus
1. Particulate matter collected on a glass fiber filter is ignited at low
temperature to destroy organic matter. The ignited filter is heated with
dilute HCl, which extracts the phosphorus and converts it to orthophosphate. The phosphorus is then analyzed by a version of the reactive
phosphorus method.
a. 1 N HCl – 82.64ml of 12.1 N HCl made to 1000 ml with DI (you will
need some for flushing the spec also)
b. Ammonium Molybdate
• In 500ml volumetric add 15g of ammonium paramolybdate,
(NH4)6Mo7O24 *4H2O and approximately 350ml DI water, shake/stir
till dissolved, make to volume (500ml) with DI and transfer to plastic
amber bottle (transparent bottle can be used, just keep out of
sunlight).
c. 4.8N Sulfuric Acid (H2SO4)
• In ice bath Add 140ml of 36N H2SO4 to 900ml DI water. Allow
solution to cool and store in a glass bottle.
d. Ascorbic Acid Solution
• In 500ml volumetric add 27g ascorbic acid and approx. 350ml DI
water, shake/stir till dissolved, make to volume with DI (500ml).
Store in plastic bottle in freezer. Thaw for use but refreeze. Stable
for many months but should not be left at room temperature. Tip:
separate solution into plastic disposable scint vials or similar
containers, in this way you can thaw out only the amount needed.
e. Potassium Antimonyl Tartrate (careful, toxic)
• 0.34g potassium antimonyl tartrate, fill to volume with 250ml DI
water, warm if necessary. Store in glass or plastic, stable for many
months.
f. Mixed Reagent 500ml (have samples ready, make daily, only keeps 68 hours)
• Final amount can be modified depending on number of samples.
Reagent should be made from the following proportions in the order
given.
Volume
100ml
Reagent
Ammonium
Molybdate
4.8N Sulfuric Acid
Comment
100ml
Ascorbic Acid
Solution
Should turn pale
yellow color
50ml
Potassium
Antimonyl
Tartrate
250ml
43 |
Arctic LTER Streams Protocol
1. PROCEDURE
a. Dry filters in petri dishes (with the lids slightly open at 70º C for 1 hour,
I typically dry the blank and standard filters in bulk in tin foil.
b. Prepare standards and blanks by placing ashed GF/F filters in marked
reaction tubes and adding appropriate amount of primary stock
standard (see below). Stock is added directly to blank/std tubes which
contain filters.
• The following is a table of amount of primary (1000uM) stock to
add, and µmols of PO4 in each tube after standard is added:
1º Std
(µl)
0
25
50
100
150
200
•
µmols
PO4
0.0
0.025
0.050
0.10
0.15
0.20
Sometimes the filters don’t absorb all of the liquid very well – try to
put it directly onto the filter and let it sit for a while in the drying
oven. If it doesn’t get fully absorbed make a note.
b.
c.
d.
e.
f.
Cover each tube with foil
Place tubes in muffle furnace at 500º C for 1 hour
After sample has cooled, add 2 ml of 1 N HCl and 10 ml of DI, Vortex
Cap samples tightly and place in oven at 104º C for 2 hours
After samples have cooled, add 2.5 ml of mixed reagent (see above) to
each tube, Vortex
g. Re-cap and place tubes in the dark
h. Allow 30 minutes for color reaction to occur, std should be noticeably
blue, and read standards and samples at 885 nm on a
spectrophotometer within 2 hours.
i. Obtain a regression equation y=mx+b where y is the absorbance, m is
the slope of the linear regression, x is the µmols of PO4, and b is the
intercept. Calculate the amount of PPO4 in each sample tube (in
µmols) using this standard curve:
µmols = (y - b) / m
The concentration of PPO4 in the water sample (in µmols/L) is then
determined by dividing the µmols by the amount of water filtered for
each sample (in liters).
44 |
Arctic LTER Streams Protocol
E. CHN
These samples are analyzed in the Jeffords Building at UVM with a CN auto
analyzer.
The FlashEANC Soil Analyzer is calibrated with aspartic acid, and uses glycine
as a secondary QC. It uses a combustion method to convert the sample
elements to the simple gases (CO2, H2O, and N2). The sample is first oxidized in
a pure oxygen environment; the resulting gases are then controlled to exact
conditions of pressure, temperature, and volume. Finally, the product gases are
separated. Then, under steady-state conditions, the gases are measured as a
function of thermal conductivity. A known standard is first analyzed to calibrate
the analyzer in micrograms. The calibration factor is then used to determine
unknowns. All quantitation is performed on a weight percent basis, using a
gravimetric technique. The system uses a steady state, wavefront
chromatographic approach to separate the measured gases. This approach
involves separating a continuous homogenized mixture of gases through a
chromatographic column. As the gases elute, each gas separates as a steady
state step, with each subsequent gas added to the previous one. Consequently,
each step becomes the reference for the subsequent signal.
It is good to have some filters pelletized before beginning so that you can run
samples at the same time as pelletizing. Pelletize standard K factors and blanks
first.
1. On the day before the filters are to be pelletized, place them in a drying
oven at 37C for at least 12 hours. Then transfer the dried filters to a
dessicator. Pull a vacuum on the dessicator if possible.
2. On the day of use sign into the log book
3. Check the pressure on the helium (20 psi) and oxygen (16 psi) tanks.
4. Check that the printer is on. Check the carousel autosampler and set it to
position 1
a. make sure the printer paper is loaded properly and that it will not roll
over on itself (pull it forward so that it will feed to the floor)
5. Turn the gas saver off – press Parameter key and type 22. Press 2 for off.
6. Purge by pressing the PURGE GAS key. Purge helium for 100 seconrds
and Oxygen for 30 seconds.
7. Check the settings to make sure it is set for filters
a. Parameters 29
b. Water concentration – yes
c. Filter PPM
d. Filter volume – unless all samples were the same volume input
1000mL – can process data to correct for actual amount filtered
8. Check to determine how many more samples can be run on the reduction
column and the filter collector – don’t plan on running more than it is set
for
9. Take the filters to the instrument room. Clean the working space with
ethanol. Get the tin disks and forceps from the drawers beneath the CHN
45 |
Arctic LTER Streams Protocol
analyzer and the balance. Clean the forceps thoroughly with kimwipes and
ethanol. Use the forceps to handle the tin disks and the filters; do not
use your fingers!
10. To pelletize a filter, center the filter on a tin disk, sample side up. Make
sure you have only one tin disk; they're very thin and it's easy to pick up
more than one at once. With the forceps, fold the disk in half so that the
filter is on the inside. Fold the disk again lengthwise (you should now have
a long thin strip rather than a quarter-circle; see the manual).
11. Fold the ends of the strip over, and then roll the strip into a sausage or
jelly roll shape. If you roll it up too tightly, you may rip the tin, so be careful.
When it is rolled up, no part of the filter inside should be visible.
12. Place the "sausage" (jelly roll if you're vegetarian) into the pelletizing
cylinder. Put the cylinder in the widemouth-side of the stage mount. Pull
down the handle part way and line up the press with the opening of the
cylinder. When they are lined up, pull the handle all the way down and
press the sausage.
13. Lift the handle and remove the cylinder. Flip the stage mount over so that
the narrow-mouth side is up. Put the cylinder back on the stage, line up
the press and the opening, and pull the handle down. The press will knock
the pellet into the narrow-mouth opening.
14. Using the forceps, remove the pellet and place it into a pellet-holding tray
(there are several trays in the instrument room). It is essential that you
keep track of which pellet is in which hole in the tray. The holes in the
trays are marked (A1, A2, A3, etc.), so use the datasheets provided (titled
"AUTO RUN Sample Information (Filters)") to record which pelletized
sample is in which hole. Leave the first row (A1-A12) empty; you will put
your initial standards and blanks in that row. About every 10 pellets, skip
two holes (a standard blank and a filter blank will go in these holes).
15. Calibrate the microbalance in the corner, using the instructions provided in
the booklet on top of the balance. The calibration weights are in the
drawer under the balance. The range setting should be 20 mg.
16. Pack standard blanks (also called a K factors):
a. Place an ashed 25-mm GF/F filter on a tin disk, fold them in half with
the forceps, and then unfold partially so that there is a crease down the
middle of the filter and tin.
b. Place a counterweight pellet on the left pan of the balance (there are
several counterweights in a small plastic box; however, some were
made with more than one tin disk and will cause an error on the scale,
so keep trying until you get the counterweight with only one tin disk).
c. Place the creased filter and disk on the right pan of the balance, lower
the pan arrest, and zero the balance. Raise the pan arrest.
d. Using the spatula, place a small amount of acetanilide on the creased
filter. Lower the pan arrest and check the weight. Add or remove
acetanilide until you have between 2 and 3 mg of acetanilide. Record
the weight on a Preliminary Standardization Data Sheet (available in
folder on table in the CHN room).
46 |
Arctic LTER Streams Protocol
e. VERY CAREFULLY, using the two pairs of forceps, roll up the disk
into a sausage as described above. Make sure you don't spill any of
the acetanilide and that you don't rip the tin. Start over if you do either
of these things, or else the weight in the next step will not be correct.
f. Pelletize the sausage and reweigh. If the weight is greater or less than
the original weight by more than about 0.010 mg, start over. If not,
then record the second weight.
g. Still using the forceps, place the pellet in a designated hole in the pellet
tray. The first five K factor pellets will go on row A (in holes A1, A3, A5,
A7, and A9). Subsequent K factor pellets will go within the samples; as
you recall, every ten samples or so, you left two holes blank. The first
of these holes is for a K factor. So, place the K factor pellet in the first
of a pair of empty holes and record the hole number and the weight of
the acetanilide in the K factor pellet on the data sheet (use the
Preliminary Data Sheet if the pellets are going into row A and the Auto
Run Sample Info data sheet if the pellets are being implanted within
the samples).
17. Pack filter blanks:
a. Using the forceps, place a blank 25-mm GF/F filter that you brought
back from the field onto one of the tin disks.
b. Pelletize the blank filter in the same manner as the sample filters
(described above); do not add acetanilide.
c. Transfer the pelletized blank filter to the pellet tray. Place in one of the
empty holes designated for blanks. The first four blanks will go on row
A (in holes A2, A4, A6, and A8). Subsequent blanks will go within the
samples; as you recall, every ten samples or so, you left two holes
blank. The second of these holes is for a blank. So, place the blank
pellet in the second of a pair of empty holes and record the hole
number on the data sheet (use the Preliminary Data Sheet if the pellets
are going into row A and the Auto Run Sample Info data sheet if the
pellets are being implanted within the samples).
18. At this point, you are ready to run your samples. If, however, the CHN
machine is already in use or not ready to run, tape your pellet trays shut,
label them, and place them in a dessicator with activated dessicant and
draw a vacuum on them until the machine is available.
19. When the machine is available, check with Marshall Otter to make sure
that there is enough helium, oxygen, and nitrogen gas, and that the
columns are fresh enough to run all of your samples.
20. When you are ready to start a run begin with Kfactors and blanks to
calibrate the machine.
21. Run an unweighed conditioning sample of acetanilide as a single run
sample
a. use 2-3mg acetanilide
b. press SINGLE RUN key
c. when prompted by 1 BLANK 2 KFACT 3 SAMP type 3 to select sample
d. when promted, type in sample ID and estimated weight
47 |
Arctic LTER Streams Protocol
e. press START to begin
22. Next, run 3 oxygen blanks
a. press SINGLE RUN key
b. select 1 for blanks
c. when prompted for the number of blanks type 3
d. press START to begin
e. Blank values should fall within these ranges
• C blank = 10-50 +/- 30
• H blank = 100-300 +/- 100
• N Blank = 50-120 +/- 16
23. After blank runs, alternate between K-factors and filter blanks
a. run using the SINGLE RUN key
• for K-factors select 2
° Acetanilide is type S1
° type in the second weight of the K-factor
° press START to begin
b. If K-factor varies more than a 0.5% from the default or previous
running mean an error will appear. If they are not ok additional Kfactors and blanks should be run
24. Once the running mean for K-factors is acceptable, you can begin running
samples
a. A K-factor and filter blank should be run after every 10 samples
• If the K-factor is out of tolerance after samples, run more K-factors
and filter blanks
25. For your samples you can use the AUTO RUN
a. Place your samples in the holes in the carousel – lining up the
numbers with your datasheet so that you know what each sample is
b. Press AUTO RUN
c. begin typing in the names of the sample
d. when you get to a Kfactor remember to type it in as a S1 kfactor and
enter the weight
26. You can begin running and continue to add to the AUTO RUN as you
complete packing more samples
27. Keep an eye on the machine – make sure that it isn’t going out of
tolerance because that can indicate that the reduction column is done and
you can lose samples, also make sure that the carousel is moving
properly and the pellets aren’t getting caught when it turns
28. Errors – check the manual to help determine issues and how to fix it
29. Shut down
a. fill out log book
b. turn gas saver ON
c. set time and date
d. Make sure CHN is left in Standby
e. clean up
48 |
Arctic LTER Streams Protocol
F. Anions
These samples are analyzed in the laboratory in Jeffords Hall on campus with
the ion chromatograph. Samples pass through a pressurized chromatographic
column where anions are absorbed by column constituents. As the eluent, a
liquid that extracts ions, runs through the column, ions begin separating from the
column. Concentrations of certain ions (such as Cl-1 and SO4) are determined by
their retention time as they are separated from the column.
1. Materials
a. Dionex chromatographer
b. Stock solutions for Cl-1 and SO42
c. 7 100-ml volumetric flasks
d. Nanopure water
e. 100- and 1000-l pipettes (calibrated), tips
f. 7 125 ml poly bottles to store standards
2. If Total Anion concentration is < 600 um the Dionex can be set up to run
on the Fast column (~ 3 min / sample). Make sure that the Dionex has
plenty of regenerant and eluent.
a. Regenerant (2 liters)
• 100 ml .5 N H2SO4 (27.7 ml H2SO4 in 2 liter DI)
• Fill to 2 Liters with DI
b. Eluent (2 liters) Should be relatively fresh.
• .4240g Na2CO3 and
• 0.0252 g NaHCO3
• Fill to 2 liters
3. Standard stock solutions for chloride and sulfate can be made in the same
volumetric
a. Chloride = Sodium Chloride NaCl (FW = 58.44)
• 10 mM NaCl = .05844 g / 100 ml
b. Sulfate = Sodium Sulfate Na2SO4 (FW = 142.04)
• 10 mM Na2SO4 = .142. g / 100ml
4. Standards:
a. 2 uM Cloride & sulfate = 20 ul 10 mM NaCl & 20 ul 10 mM Na2SO4
/100 ml DI
b. 5 uM Cloride & sulfate = 50 ul 10 mM NaCl & 50 ul 10 mM Na2SO4
/100 ml DI
c. 10 uM Cloride & sulfate = 100 ul 10 mM NaCl & 100 ul 10 mM Na2SO4
/100 ml DI
d. 15 uM Cloride & sulfate = 150 ul 10 mM NaCl & 150 ul 10 mM Na2SO4
/100 ml DI
49 |
Arctic LTER Streams Protocol
e. 30 uM Cloride & sulfate = 300 ul 10 mM NaCl & 300 ul 10 mM Na2SO4
/100 ml DI
5. The attenuation will need to be changed for the different concentrations in
the standard curve. On the fast column the following work well. For
actual area under the curve multiply the attenuation by the area.
Standard concentration
(uM)
1
2
2
5
3
10
4
15
5
30
attenuation
1
1
3
3
10
6. To run a sample:
a.
b.
c.
d.
50 |
Inject sample into port (>1 ml)
Hit [Load/ Inject] (valve will switch)
Change attenuation if needed
Plot sample [Shift/Down] [Plot] [Enter]
Arctic LTER Streams Protocol
G. Cations
Rains, Theodore, C. 1984. Atomic Absorption Spectrometry. Water
Analysis, Vol II Edited by Roger A. Minear and Lawrence H. Keith.
These samples are analyzed in the laboratory in Jeffords on campus with an
Absorption/ Emission Spectrophotometer.
1. Materials:
a. Reference standard solutions for atomic absorption + 1%:
b. Na+, (1000 ppm 100 ml poly bottle Fisher cat # SS 139-100)
c. K+, (1000 ppm 100 ml poly bottle Fisher cat # SP 351-100)
d. Mg+2, (1000 ppm 100 ml poly bottle Fisher cat # SM 51-100)
e. Ca+2 (1000 ppm 100 ml poly bottle Fisher cat # SC 191-100)
f. 20 100-ml volumetric flasks (5 per element)
g. 5-ml volumetric pipette & pipette bulb
h. 1000-ul pipette (calibrated), several tips for dilution’s
i. deionized water
j. HCl (trace metal grade)
k. 10% Lanthanum Cloride (LaCL3 FW 371.38) (for calcium)
l. Absorption/Emission spectrophotometer
m. clean, dry scintillation vials (at least one per sample)
n. 10 ml Re-pipette
o. Samples should be preserved by acidifying to ~ pH 2 (100 ul Ultrex
HCL / 60 ml sample.
p. Sign up for use of the Absorption/Emission Spectrophotometer in the
Instrument Room
q. The week that you plan to run your samples, make standards using
stock solutions kept in the back room of the Aquatics Lab, second
floor, Ecosystems. For each ion, make standards in concentrations of
1, 2, 3, 4, and 5 parts per million (ppm). Make the standards in 100-ml
volumetric flasks (labeled) under the hood. *Use trace metal grade
HCl, measured in the volumetric pipette.
•
•
•
•
•
1.00 ppm = 100 ul stock, 5 ml HCl, water to fill line.
2.00 ppm = 200 ul stock, 5 ml HCl, water to fill line.
3.00 ppm = 300 ul stock, 5 ml HCl, water to fill line.
4.00 ppm = 400 ul stock, 5 ml HCl, water to fill line.
5.00 ppm = 500 ul stock, 5 ml HCl, water to fill line.
IMPORTANT NOTE:
1. Chemical interference in flame AAS originates during dissociation of the
analyte in the flame cell forming a compound that volatilizes at a different rate
than the standard. One method of alleviating this type of interference is
51 |
Arctic LTER Streams Protocol
through the addition of a releasing agent or protective chelate. A releasing
agent is a metal or salt that forms a more stable compound with the
interferent than the analyte. For calcium analysis the addition of lanthanum,
which acts as a releasing agent and reduces interference with the calcium
peak from Al, Si, PO43-, and SO42- is required.
1.1. Lanthanum Cloride (LaCL3 FW 371.38) should be added to a 1% final
concentration to all samples and standards. 10% LaCL3 = 37.13 g / liter.
1.2. For each calcium standard, add 10 ml of 10% lanthanum solution
1.2.1. 1.00 ppm = 100 ul stock, 5 ml HCl, 10 ml of 10% LaCL3 water to fill
line.
1.2.2. 2.00 ppm = 200 ul stock, 5 ml HCl, 10 ml of 10% LaCL3 water to
fill line.
1.2.3. 3.00 ppm = 300 ul stock, 5 ml HCl, 10 ml of 10% LaCL3 water to
fill line.
1.2.4. 4.00 ppm = 400 ul stock, 5 ml HCl, 10 ml of 10% LaCL3 water to
fill line.
1.2.5. 5.00 ppm = 500 ul stock, 5 ml HCl, 10 ml of 10% LaCL3 water to
fill line.
1.3. For each 1 ml of sample add one of the following depending on
concentration.
1.3.1. 1 ml of (200 ml 10% LaCL3 in 1 liter DI) multiply reading by 2 for
final conc.
1.3.2. 9 ml of (Add 100 ml 10% LaCL3 to 900 ml DI) multiply reading by 10
for final conc.
2. Fill the large beaker (inverted on the front of the spec) with DI water (in the
carboy on the shelf to the left of the gas cylinders). Turn on the exhaust fan
(switch by the door).
3. When it is time to run the samples on the Absorption/Emission spec, fill out
the log book in the Instrument Room (on the little table next to the AAS. Start
up procedure varies depending on the elements being run. Emission
methods are used for K+, Na+ analysis, while absorption methods are used for
Mg+2 , Ca+2. For absorption you will need to fill out the log book for lamp and
Continuous Operating current (Mg+2 , Ca+2. 15 ma).
4. Turn on the air (the knob is on the wall above the gas cylinders). Then, turn
on the acetylene. The low stage should stay above 15 (in red), and the high
stage should not fall below 70 psi. If a new tank is needed, call Martha @
7272).
5. Initial set up for all elements.
5.1. Push [Power on].
5.2. Make the following control settings:
5.2.1. SIGNAL - LAMP
5.2.2. GAIN - Fully counterclockwise
5.2.3. BG Corrector - AA
52 |
Arctic LTER Streams Protocol
5.2.4. LAMP current control - fully counterclockwise
5.3. Set the SIGNAL control to set up. Set the SLIT control to the setting
appropriate for the element of interest. Adjust with the course adjust
wavelength control to obtain the wavelength of interest.
5.3.1. K+, wavelength = 766.5 and slit width = 0.7.
5.3.2. Na+, wavelength = 589.0 and slit width = 0.2.
5.3.3. Mg+2 wavelength 285.2 and slit width .7
5.3.4. Ca+2 wavelength is 422.2 and slit width is .7)
5.4. Push [Flame on]. If an error message appears (E 50) it is due to air in
the lines, push [Flame on] again.
5.5. Put the sipper tube into the DI in the big beaker.
5.6. Adjust the ratio of air/fuel so that it is about 18:40.
6. Emission methods (use for K+, Na+ analysis)
6.1. If a lamp is installed be sure to unplug it at this point.
6.2. Turn the [Signal] knob to EM (emission)
6.3. While aspirating a blank solution press [AZ] to auto-zero the spec.
6.4. Aspirate the most concentrated standard solution, optimize the
wavelength by adjusting the [Fine adjust] dial slowly back and forth to provide
maximum energy. Adjust the gain control as necessary to keep the LAMP
/ENERGY display on scale.
6.5. Pick two standards between which samples fall (range of 1 ppm). While
sipping the higher of the two standards adjust GAIN to provide a reading of
about 75 on the LAMP/ENERGY display. Note that every time standards are
changed the gain must also be changed.
6.5.1. NOTE: Calibration curves for flame emission measurements may
not be accurate over extended concentration ranges
6.6. Change the integration time to 3 seconds by hitting 3 [t],
6.7. Set values for the standards: hit 1.00 and [s1] and 2.00 [s2] for 1ppm
and 2 ppm if two decimal places are desired use two.
6.8. Hit [az] to auto zero while sipping DI. Sip low standard and wait for
absorbance to stabilize and hit [s1] repeat for standard 2
6.9. Sip s1 again to check reading if not similar redo calibration
6.10.Run all standards in a given range while periodically checking high
standard to be sure gain has not dropped if it has either edge it back or reset
gain to 75.
7. Absorption (Mg+2 , Ca+2)
7.1. Place correct hollow cathode lamp in lamp holder and plug in.
7.2. Turn the lamp current control until the LAMP/ENERGY display shows
the proper lamp current, as given on the lamp label for continuous operation
(continuous operating current for Ca+2 & Mg+2 lamp is 15).
7.3. Turn signal knob to set up. Turn the FINE ADJUST wavelength control
slowly to obtain a maximum reading on the LAMP/ENERGY display. Use the
53 |
Arctic LTER Streams Protocol
gain control to adjust the maximum reading to 75. If the gain should become
too high, an over range reading of EE will be obtained.
7.4. Align lamp by turning the two alignment knobs on the lamp holder to
maximize the LAMP /ENERGY display reading. Again use the gain control to
make the maximum reading 75.
7.4.1. NOTE: As the lamp warms up, the display value may increase
slightly drop gain back down to final setting of 75
7.5. Close the lamp compartment door.
7.6. Wait ten minutes
7.7. Set the SIGNAL control to concentration.
7.8. Hit [az] to auto zero while sipping DI. Sip low standard and wait for
absorbance to stabilize and hit [s1] repeat for standard 2
7.9. Sip s1 again to check reading if not similar redo calibration
7.10. Run all standards in a given range while periodically checking gain and
readjust to 75 as lamp warms
8. Shut down
8.1. after last sample let run for 10 minutes
8.2. turn signal, gain, lamp counter clockwise
8.3. fill out log book
8.4. turn flame off
8.5. pull sipper out of DI and dump DI
8.6. shut off air and acetylene
8.7. empty lines of air and acetylene by hitting [check 0-2 ] [check fuel]
8.8. turn off machine
8.9. turn off fan
8.10.unplug lamp
54 |
Arctic LTER Streams Protocol
H. Alkalinity
GRAN TITRATION
Principle:
Alkalinity is the measurement of the Acid Neutralizing Capacity (ANC) of a
water sample. Alkalinity is usually reported in units of milliequivalents per
liter of sample (meq/L). In Toolik area waters, ANC is due primarily to
HCO3, CO3-2, OH-, and certain organic bases. Of these, HCO3 is usually
by far the most important species. We are interested in measuring
alkalinity for a couple of reasons. When coupled with a measurement of
pH, alkalinity can be used to compute total dissolved inorganic carbon
(needed for primary production measurements), and the partial pressure
CO2 gas in the water (useful for atmosphere-water interaction studies). In
waters which have a near-neutral pH (most of the Toolik area), alkalinity
correlates well with the total concentration of dissolved ions, and hence
can be useful in categorizing the overall ionic state of a water sample.
Also, when used as a long-term monitoring tool, it can detect acidic
impacts on lakes and rivers. Alkalinity is usually measured by titrating a
water sample with a strong acid. The alkalinity is a measurement of the
amount (equivalents) of acid needed to exactly neutralize the original ANC
of the water sample. There are two common methods of performing this
titration; both are based on monitoring pH as acid is added to a water
sample. The procedure outlined below is based on the “Gran”
methodology. In the Gran method, a series of pH measurements are
made. The alkalinity is determined by an analysis of the rate at which pH
changes in response to acid additions. In practice, the alkalinity is
computed with a computer program written for this purpose.
Collection and storage of water samples:
Either plastic or glass sample bottles are acceptable. If sample bottles
were previously acid-cleaned, then bottle should be rinsed thoroughly with
sample water if possible to insure that sample is not exposed to acid.
Samples CANNOT be treated with either acidic or basic preservatives.
This includes all acids and bases, and many preservatives such as
formaldehyde. Samples should not be frozen. Analyses should be
conducted within 24 hours. EITHER filtered or unfiltered water may be
used for analyses.
EXCEPTIONS:
1. If there is going to be a delay of more than one day between collection
and analysis, then filtering is recommended. Alkalinity can change if
biological activity results in either the production or dissolution of organic
particles. If samples are going to be kept for more extended periods, then
they should be stored in completely full, tightly stopper bottles.
2. If water samples are suspected of containing particles of calcium
carbonate, sample MUST be filtered before analysis. If the sample is
55 |
Arctic LTER Streams Protocol
turbid, but composition of particles is not known, then filtering is an
advisable precaution.
3. Anoxic water samples present special problems. Anoxic water samples
from the Toolik area usually contain high concentrations of dissolved iron.
When the anoxic water comes in contact with oxygen from the
atmosphere, the dissolved iron will begin to precipitate. Iron precipitation
strongly decreases alkalinity, and must be avoided. The best solution is to
collect anoxic water samples by completely filling to overflowing GLASS
dissolved oxygen bottles. Bottles must remain tightly stoppered until
analysis. Bottles should be opened individually JUST BEFORE analysis is
to begin.
pH calibration for the Mettler Auto-Titrator:
1. Turn on the Mettler Auto-Titrator, switch is on the back located on the
lower left corner .
2. Remove the gray cap covering the opening of the pH tube, near the top of
the pH tube.
3. Press "reset".
4. Press "1" then "Elec Calib".
5. On the screen you will see a small red light beside buffer A.
6. Fill a Mettler cup with approximately 50-ml of 4.0 buffer solution.
7. Screw the Mettler titration cup in place.
8. Press "start", the stirrer should turn on if not check that the stirrer knob is
set to 3.
9. Once the Mettler has calibrated the 4.0 buffer, the red light will light up
beside the buffer B on the screen.
10. Empty the 4.0 buffer back into its container, rinse with DI and wipe dry the
Mettler cup, and fill with the approximately 50ml of 7.0 buffer solution.
Screw the Mettler cup in place.
11. Press "start", the stirrer should turn on.
12. The Mettler Auto-Titrator will beep once it is done calibrating and usually
"0" will appear on the screen once it is ready.
13. Pour the 7 buffer solution back into its container, rinse and dry the Mettler
cup.
Prior to running samples:
1. All samples should be at room temperature when analyzed because
pH is strongly influenced by changes in temperature.
2. The laboratory room where the measurements are made must be free
of acid and base fumes. These vapors will dissolve in water and
change the alkalinity. Likewise, if samples are filtered or otherwise
processed in the laboratory, care must be taken to avoid
contamination.
3. The methodology described below presumes that the alkalinity
analyses will be made using the Mettler “Auto-Titrator.” The pH
electrodes should be calibrated at least once per day as described in
56 |
Arctic LTER Streams Protocol
pH calibration section above, usually when the Mettler titrator is turned
on. The normality of the acid used for titration MUST be known very
accurately. The acid routinely used at Toolik is 0.1 N H2SO4, and is
usually purchased commercially. However, other normalities may be
used if necessary.
4. Before beginning analyses, make sure acid dispensing and delivery
tubes on the titrator are free of air bubbles. This can be a common
problem at the start of each day and after the burette is refilled.
Bubbles can usually be removed by gentle tapping to free the bubble
from the tubing walls, followed by the dispensing of several mL of acid
to flush the bubble from the tube. The dispensing of acid is done my
entering a value, e.g., 2 or 5 and pressing the "dose ml" button.
1.
2.
3.
4.
5.
57 |
Running the alkalinity titration
Between samples, rinse the electrode 10 x with the DI pump and then wipe
dry the pH electrode, stirrer and Acid tube with a kim wipe to remove all water
droplets.
Special Mettler titration cups are used for the samples. Titration cups MUST
be clean and completely dry before use. Fill the titration cups with a KNOWN
volume of sample. 50-mL samples are routinely used for Toolik
measurements; however, other volumes may be used. The accuracy of the
alkalinity measurement depends DIRECTLY on the accuracy of the sample
volume. The preferred method for measuring and dispensing sample
volumes is a volumetric pipette. The second choice for measuring and
dispensing samples is a graduated cylinder. Make sure to rinse the pipette
with DI or sample rinse between samples.
Screw the titration cup into the electrode/stirrer/dispenser holder.
Turn on the stirrer, knob on left side of Mettler, and set to 3.
Begin alkalinity titration by pressing the following keys on the Mettler:
5.1. Press “pH meas” , the stirrer will turn on and watch the pH rise to 7-8.
Occasionally the samples pH will not reach 7 in which case continue to
the next step, but wait at least 2 minutes.
5.2. Press “Reset”
5.3. Press “ENDPOINT” to set pH endpoint for initial titration. Enter a value of
4.1 and press "ENDPOINT” again. If the Mettler is already set to 4.1, you
need only press "ENDPOINT" once.
5.4. Press “START” four times to begin titration. MAKE SURE stirrer is on
and is at a moderate speed (about 3).
5.5. Titrator will beep when the endpoint has been reached.
5.6. *Write down amount of acid added*
5.7. To obtain the pH of the sample once the endpoint has been reached,
press "pH meas", stirrer should come on. Allow sample to stir for at least
one minute. This allows CO2 to degas from the sample and permits
stable pH measurement. Once the pH stabilizes write down the pH, if you
have difficulty in determining the pH with the stirrer on, you can turn off
the stirrer by turning the knob and write down the pH value before the
Arctic LTER Streams Protocol
values start to increase. The pH values will start to increase as soon as
the stirrer is turned off.
5.8. At this step in the analysis, pH should be between 3.95 and 4.05. If pH is
>4.05, reset ENDPOINT for 4.0 and proceed again as above. When
proper pH range has been achieved, record BOTH mL and pH.
5.9. Press “RESET,” enter 0.02, and press “DOSE ml.” This will cause 0.02
mL of acid to be dispensed into the titration cup.
5.10.Press "pH meas", the stirrer will turn on. After the stirrer has been on for
1 minute or once the pH stabilizes write down the pH, or you can turn off the
stirrer by turning the knob and write down the pH value before the values start
to increase. Record BOTH pH AND the TOTAL volume of acid, which has
been added to that point.
5.11.Press “DOSE mL” to dispense more acid and repeat as above. A pH
change of about 0.1 pH unit is desirable. It will probably be necessary to
increase the amount of acid dispensed as the titration proceeds such as using
0.02 ml, 0.03 ml, 0.04 ml, 0.05 ml, 0.06 ml, 0.07ml, if you are close to a pH of
3.1 and still need to add acid to get 10 points you can add smaller amounts
like 0.01ml..
5.12.Continue until at least eight readings have been determined which cover
the pH range of approximately 3.0 - 4.0. Again, it is necessary to record
BOTH the pH and TOTAL volume of acid dispensed for each set of
measurements.
6. Compute alkalinity by entering data into Quattro Pro spreadsheet Alkalinity
program. Data entered for each sample are 1) sample volume; 2) acidic
normality; 3) sets of volume and pH measurements. Alkalinity is computed by
running the macro program which is part of the spreadsheet. Check the
output file for Correlation Coefficient and Predicted Acid Normality. If
Correlation Coefficient is not greater than 0.98, or if Predicted Acid Normality
differs greatly from actual acid normality, then there has probably been an
error in the titration or data entry.
Alkalinity Calculation on Spreadsheet:
Example of data entry in spread sheet:
Sample Hershey Cr. St.4,
500m
Date
23-Jul-01
collected
Date run 26-Jul-01
Acid N
0.1005
Sample
0.05
V (ml)
Acid (ml) PH
[H]
0.3471
3.994 0.000101
0.3671
3.867 0.000136
0.3871
3.771 0.000169
58 |
Arctic LTER Streams Protocol
0.4071
0.4371
0.4771
0.5171
0.5671
0.6271
0.6971
3.69
3.585
3.484
3.395
3.31
3.224
3.14
0.000204
0.00026
0.000328
0.000403
0.00049
0.000597
0.000724
END0.000291
POINT
(L)
R^2
0.999939
ALK
0.585914
(meq/L)
[H]: 10 ^ (-1*(acid at 4.1 endpoint)
E.g., 10^(-1*0.3471) = 0.000101
END-POINT: intercept (all acid points, [H] points) / 1000
E.g., intercept (0.3471…0.6271,0.000101…0.000724)/1000 =
0.000291
ALKALINITY: [(endpoint*acid N) / sample volume] * 1000
E.g., [(0.000291*0.10005) / 0.05]* 1000 =0.585914
Comments:
1. Do not let the pH probe sit in the air, if you will not be running samples for a
prolonged period, screw in place a Mettler cup filled with 60 ml of DI water.
2. When finished using the Mittler Auto-titrator remember to place the gray cap
back onto the pH probe
3. Turn off the Mettler Auto-titrator at the end of the day.
4. Occasionally the screw that holds the pump into place may become loose and
thus the Mettler Auto-titrator will be titrating an incorrect amount of acid.
Check that the amount of acid titrated is the amount entered, if it isn't check
the screw under the pump platform.
5. RUNNING STANDARD
5.1. run a known alkalinity standard to test the acid used. Run these
standards whenever a new acid is used.
IV.
Appendix A – Formerly Used Methods
A.
2x2 rock scrubs
1. The following methods are based on methods from the LTER
database, previous protocol sheets, and Peterson et al. (1993), and
have been modified into outline form.
59 |
Arctic LTER Streams Protocol
2. Materials
3. At each station, rinse the wash basin, scrub brush, slide holders, and
wash bottle 3X with river water. Fill the wash bottle with river water.
4. From a riffle, select 5 (or 3 if only doing 3 reps) rocks that fit the
following criteria:
a. rocks with no filamentous algae or moss (to eliminate
overestimates of chl due to filamentous algae or moss)
b. rocks with fairly smooth upper surface (uneven surfaces prevent
efficient removal of epilithon)
c. rocks that have been submerged for a long period of time.
d. It is possible that very few rocks at some stations will meet all of the
above criteria. If you must select rocks that do not fit any or all of
the criteria, make careful and thorough notes describing the
deviations.
5. Place the slide holder over a smooth portion of the upper surface of the
rock. With the brush, scrub the area within slide holder. Hold the rock
over the basin so that all scrubbate falls into the basin.
6. With the wash bottle, rinse the scrubbed area, the holder, and the
brush into the basin. Pour any remaining rinse water into the basin.
Record the initial slurry volume used (volume of rinse bottle).
7. Pour the contents of the basin into a labeled (by river and station)
centrifuge tube or bottle. Use the funnel to facilitate pouring.
8. Repeat steps 2-8 for each rock at each station. Only scrub one rock
per sample bottle at each station. You will end up with 3-5 replicate
slurries from each station (depending on how many reps you decide to
do).
B. Epilithic primary productivity
1. In addition to the studies involving the rock scrubs, rocks will be
incubated in special laboratory chambers to determine primary
productivity of the epilithic algal layer. The amount of photosynthesis
and respiration occurring on individual rocks will be determined during
the incubation period. These rocks may have filamentous algae; the
rock scrubs excluded such rocks when possible.
2. The methods below are based on methods written by Breck Bowden,
the PI for this project, in the document files of the Arctic LTER
database in 1989 and 1990 (filenames 89BOMETA.DOC and
90BOMETA.DOC). They have been modified into outline form.
a. Materials:
• carboy (optional)
• coolers
3. At the scheduled time (determined at the beginning of the field
season), collect rocks at random locations within pools or riffles at
60 |
Arctic LTER Streams Protocol
each sample station (locations vary from year to year). Select surface
rocks with a "typical" development of epilithon, based on visual
inspection of the station, for study. Reject rocks with heavy moss
growth and instead choose rocks of a uniform size and shape, that
would fit neatly in the chamber bottom.
a. In the Kuparuk River, 3 to 4 rocks of a modal shape and size
essentially covers the chamber bottom in a single layer, at a
surface density similar to that found in the river. Substrate size in
Oksrukuyik Creek is substantially smaller than in the Kuparuk
River; thus, use 4 to 7 rocks from Oksrukuyik Creek to cover each
chamber bottom. (Substrate sizes for New Reach streams will be
determined at the beginning of the summer.)
b. Collect water for the incubations in carboys (optional; Toolik Lake
water may be used instead).
c. Place rocks collected from the river in coolers, without water, to
keep them cool and moist. (If the rocks are kept submerged in
water in the coolers, delicate epilithic material will be dislodged
during transport from the field to the lab. Without water, the rocks
and epilithon can be transported with the epilithon essentially intact,
even with flocculent pool epilithon.) The time from collection and
transport to installation in a chamber is generally 1-2 h, and should
be minimized.
d. Upon return to Toolik, immediately place rocks and water into the
experimental chambers in Bowden’s polar tent.
C. Bioassays of epilithic algae
1. Bioassays of epilithic algae are done using small-scale artificial
nutrient-diffusible substrata. These bioassays produce many
replicates of numerous treatments at minimal cost and time
expenditure.
2. The methods below are based on Gibeau and Miller (1989) and have
been modified into outline form. Mike Miller is the PI for this project.
3. Materials:
a. agar vials (see below)
b. porous porcelain discs, soaked in HCl (see below)
c. wooden vial holders (see below)
61 |
Arctic LTER Streams Protocol
d. silicon sealant
e. rope to secure vial holders (should be several meters longer than
width of stream)
f. 2 metal spikes per wooden vial holder
g. pliers (bring when time to remove vials from river)
h. 125-ml urine cups (one per vial) (bring when time to remove vials
from river)
4. Three assay experiments will be conducted in each stream during
each summer. The dates and sites will be determined at the beginning
of the field season. Each experiment lasts three weeks.
5. Each experimental chamber is composed of a 10-dram plastic vial
(Dynalab Corp. #2636-0010) used as a reservoir, filled with various
nutrient-supplemented agar treatments.
6. The agar treatments are 37-ml of a 2% (w/v) Difco Ultrapure Agar
solution augmented with one each of the following treatments:
a. control (plain agar)
b. humic acid extract plus phosphorus (2 g humics/L and 0.5 ml conc.
HCl plus 0.005 moles K2PO4/L)
c. phosphorus (0.005 moles K2PO4/L)
d. ammonium (0.05 moles NH4Cl/L)
e. phosphorus plus ammonium (0.005 moles K2PO4/L + 0.05 moles
NH4Cl/L)
f. vitamins (B1 0.1 mg/L, plus Biotin 5 mg/L)
g. a trace metal mixture (Woods Hole formula plus 0.0999g NTA/500
ml as a chelator; Stein 1973).
7. All agar treatments should be autoclaved to ensure sterility. (Does
this affect the vitamin treatment?)
8. The chamber is sealed with a coarse, porous porcelain or fused silica
(2.6-cm diameter) disk (crucible cover) (Leco Corp. #528-041) that
has been cleaned by soaking in a 10% HCl for 48 hr and rinsing
copiously with distilled water.
9. Heat each disc on a hot plate. Seal the agar-filled vial by placing the
hot disc on top of the vial, melting the plastic at the mouth of the vial,
and molding it around the disc.
10. Turn the vial upside-down, allowing the agar mixture to solidify in
contact with the porous disc.
11. Cap and color-code finished vials according to the treatment they
contain.
12. Arrange the vials in batches of 42; each batch will contain 6 replicates
of each of the seven treatments.
13. Place each batch in a wooden holder(s), which are strips of lumber
with pre-drilled holes (3-cm diameter) and mounted on 1.2 m x 0.31 m
62 |
Arctic LTER Streams Protocol
plywood. Secure the vials into the holes with a small spot of silicon
sealant on the bottom of each vial (use as little as possible to avoid the
effects of acetic acid leaching from the sealant).
a. At each site in the stream, secure the wooden holder to the stream
bottom. Use two restraints: a rope running between opposite
shores and looped through a hole on the upstream side of the
wooden base; and two metal spikes at each end of the board,
driven through the base and into the rocky bottom, with flat rocks
placed over the stakes at each end. This should ensure that the
boards will remain stationary on the river bottom even in high flow
periods.
b. Leave the boards and the agar vials undisturbed for three weeks.
This procedure will be done three times, with one week of overlap
between experiments.
c. When an incubation period is over, unfasten the board from the
river bottom but keep it submerged. Carefully maneuver it to the
shore.
d. Remove the vials one at a time by color code and place the discs
into pre-labeled plastic 125-ml urine cups. The discs can be
removed by gently squeezing the mouth of the vial with pliers.
e. Three discs from each treatment can be placed in the cups dry;
they will be assayed for chlorophyll α biomass. The other three
should be placed in cups with about 50 ml of water and remain
completely submerged; they will be assayed for primary
productivity.
f. Return the samples to the lab and give them to Miller’s group.
D. Ammonium – Phenol method
1. This method was used 1983 -1999.
2. Ammonium Manual Chemistry
a. In order to detect the low levels of ammonium found in most
unfertilized stream reaches and lakes, it is necessary to use a
manual method at Toolik. A blue compound, indolphenol blue, is
formed by the reaction of ammonium, hypochlorite, and phenol.
The color is intensified by nitroprusside. A spectrophotometer @
630 nm with a light path capacity of 5 cm is used.
b. The protocol below was adapted by Bowden and Finlay from the
automated phenate method described in section 4500 H of
Standard Methods (1985).
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Arctic LTER Streams Protocol
3. General Comments
a. Wear gloves at all points of this procedure because it is easy to
introduce contamination from bare hands, and the reagents are
toxic.
b. Always use ammonium-free water for all reagents. Get the water
from the nanopure unit just before you use it.
c. Acid wash all glass and plasticware (including volumetric flasks and
pipettes) in the acid bath dedicated for ammonium chemistry (to
avoid contamination from other sources). Following the acid bath,
immediately rinse 3-5 times with fresh distilled water.
d. Phenol breaks down plastic so use glass to store this reagent.
e. Use dedicated glassware for each step of the procedure, including
the same containers for the same standards (especially blanks).
Do not use glassware that has been used for nitrate chemistry,
which uses ammonium as a reagent.
f. Because the reagents are toxic, place all bottles in a wash basin in
case of spill, and work in a fume hood if available.
g. Use repipetters for dispensing reagents into the sample. This
speeds things up considerably. There are 500-ml amber bottles for
this purpose. Use of repipetters also minimizes possible
contamination.
h. When making the standard dilution series, make sure the pipetters
are calibrated properly. It is important to get very accurate dilutions
when working with such low concentrations. Alternatively, you can
use volumetric pipettes for most of the dilution series.
i. Make sure waste and old reagents are stored in the properly
labeled container, including the waste that runs through the spec.
j. Whenever possible, use DI direct from the nanopure unit, without
transporting in an additional container.
4. Reagent preparation
a. Hypochlorous acid 5% NaOCl (Bleach)
• Prepare fresh for each day. 100 ml of reagent is enough for
about 100 samples and standards.
• Add 20 ml 5% NaOCl (chlorox) to a 100-ml volumetric flask.
• Add 80 ml DI water to 100-ml mark.
• Adjust pH to 6.5-7.0 with 1 N HCl. Pour a little of prepared
solution into bottle cap or petri dish and measure with pH paper.
Add a couple of ml of HCl to reagent bottle and recheck.
Repeat until color of pH paper is correct. The acidity change
will be rapid, so do not add too much acid at any given time or
you’ll have to start over.
• Chlorox should be brought up fresh each year, or perhaps
several times during the summer. It should be stored in the
refrigerator when not used. Caution: a large head space allows
chlorox to oxidate, increasing the pH.
b. Phenate (Phenol, water, NaOH)
64 |
Arctic LTER Streams Protocol
• Prepare fresh for each day. Refrigerate in amber glass container
if saving overnight.
• *Handle with care, read MSDS for Phenol prior to use*
• To a 100-ml volumetric flask, add 9.3 ml of liquid phenol (the
liquid form is easier to use than the crystal form) using a
volumetric pipette with a pipette bulb.
• Bring to 100 ml mark with fresh DI water.
• While cooling with tap water (on ice if necessary) slowly add 3.2
g NaOH and dissolve. The addition of NaOH is an exothermic
reaction, so do not stopper the flask. Excess NaOH will speed
color development but will cause a urine yellow color to develop
in all samples. Yellow color may affect blank absorbance, so try
to avoid.
c. Nitroprusside (sodium nitroferricyanide)
• Store in amber glass container for up to 1 month.
• *Handle with care, read MSDS for nitroprusside prior to use*
• Add 800 ml DI water to a 1-L volumetric flask.
• Add 3.5 g sodium nitroprusside and swirl until dissolved.
• Fill to 1 liter with DI water.
5. Standard Preparation
a. Stock A solution (prepare at the beginning of the summer)
• Acid wash 1-L volumetric flask
• Make 1 mM solution of NH4-N from (NH4)2SO4
• MW of (NH4)2SO4 = 132.14
• 2 moles of N per mole of ammonium sulfate
• 1 M NH4-N = 66.07 g of ammonium sulfate in 1 L of DI
• 1 mM NH4-N = 0.06607 g in 1 L of water. Make a note if you end
up using a slightly different amount and adjust concentration
when making up standard curve.
• Use ammonium sulfate that has been dried (in a glass vial) in the
drying oven for 12 hours before using.
• Store solution in labeled amber container in the refrigerator
b. Stock B - Secondary stock solution (prepare just prior to use)
• Acid wash the 0.5-L volumetric flask and rinse several times with
DI
• To prereact the flask, add 20 ml of DI water, 1.2 ml phenate, 1.0
ml hypochlorous acid, and 0.8 ml nitroprusside and let sit for
several hours. Swirl to clean inside. Dump reagent in waste
container and rinse with DI 3-5 times.
• Using a calibrated pipetter, or volumetric pipettes, make a 10mole secondary stock to use for preparing the dilution series
° To a 0.5-L volumetric flask, add 5 ml Stock A, and fill to the
0.5-L mark with fresh DI
° Invert several times.
c. Dilution Series
65 |
Arctic LTER Streams Protocol
• For low levels, use 4-5 standards with concentrations less than 1
Mol.
• Use dedicated 100-ml volumetric flasks that have been acid
washed and prereacted (as described for stock B preparation).
• Series should be made just prior to use, because low standards
in particular absorb ammonium very easily. DI water should be
fresh from the nanopure filter unit.
• If using pipetter, make sure each volume is calibrated.
• Try to use one type of volumetric flask for all dilutions to avoid
dilution errors.
• Recommended series:
° 0.05 mol .5ml of stock B, 99.5 ml fresh DI
° 0.1
1ml
99
° 0.2
2ml
98
° 0.4
4ml
96
° 0.6
6ml
94
° 1.0
10ml
90
° 5.0
50ml
50
6. remaining stock B solution
° blank 100ml DI water
7. Get the dedicated 50-ml centrifuge tubes still containing reagent from
the last run, shake, and dump reagent into the proper waste container.
Rinse 3-5 times with fresh DI water.
8. Pour 20 ml of standard from volumetric flask to 50-ml centrifuge tubes
dedicated for that standard concentration. Volume does not have to be
exactly 20 ml because the reagents are added in excess.
9. Use 2 tubes for each standard and 4 tubes for the blank. In 1994, 2 of
the blanks received superadditions of reagents. To these, 1.5 times
the normal amount of reagents was added. This was helpful in
determining possible sources of contamination. Absorbance should
be equal for both types of blanks and should be less than the lowest
standard.
10. Color intensity is affected by age of reagents. Thus, reagent blanks
have to be run with each set of samples and standards.
11. Procedure for sample ammonium determination
a. Reagents should be left in sample tube from previous run. Prior to
going into field, shake reagent to prereact the entire inside surface,
then dump reagent into ammonium chemistry waste container and
rinse several times with fresh DI. This should be done for standards
and blanks as well. Shake dry the tubes.
b. Return from the field with 20 ml of filtered water in each centrifuge
tube from each station.
• Add 1.2 ml phenate reagent and swirl.
• Add 1.0 ml hypochlorous acid reagent and swirl.
• Add 0.8 ml nitroprusside reagent and swirl.
66 |
Arctic LTER Streams Protocol
• There should now be about 23 ml total in each sample tube.
• At this time, you should also add the three reagents, as
described above, to the standard dilution series and to the
blanks; remember to add 1.5x the normal reagent amount to two
of the four blanks.
• Cap tightly and place samples in dark for at least 1 hour but no
more than 24 hours. (Color formation should be complete after
10 min).
12. Reading on the Spectrophotometer (Terrestrial Trailer)
a. Samples should be read immediately after reading the standards,
or else they may develop more intense color and they will no longer
be comparable to the standards.
b. Make sure Mr. Sipper is set to sip 5ml?
c. Set wavelength to 630 nm and make sure the spec is reading
absorbance.
d. Wear gloves when reading standards and samples on the spec.
Ammonium from hands could be left on sipper tube, and reagents
are harmful.
e. Make sure the waste tube enters the ammonium waste container.
f. Sip DI water and zero the reading.
g. Sip standards so that low standards are read first (start with blanks
and work way up) After each standard, sip water through and
rezero if necessary. Sip DI between high standard and first sample
to remove any residual NH4.
h. Sip again to make sure all residual material is removed and the
reading is stable. Rezero the spec.
i. Sip your samples through and record the values in a notebook.
After reading the entire sample series once, read them each a
second time. Make sure you avoid sipping air into the spec, as this
might interfere with your readings.
j. Between samples, wipe the sipper tube dry with a kimwipe so that
contamination is minimized.
k. If a sample absorbance is higher than highest standards, you will
need to dilute the sample. (can you do this in the tube?)
13. Regression
a. Subtract the average absorbance of the blanks from all standard
readings. Then regress the concentrations of the standards against
their blank-corrected absorbances. You should get an r2 of at least
three nines (e.g., r2=.9992).
b. Use the regression equation to calculate the concentration of
ammonium in the samples.
14. Notes
a. We have found it better to leave reagent in sample and standard
tubes and flasks from the last run until right before the next run.
67 |
Arctic LTER Streams Protocol
Just before run, dump reagent in waste container and rinse tubes
with fresh DI.
b. Standards should be made fresh just prior to run. Low standards
that have sat all day tend to absorb ammonium. The same holds
true for DI water. Always get fresh water from nanopure filtration
unit.
15. Checklist
a. Beginning of summer
• Make ammonium sulfate stock solution
• Make nitroprusside reagent
• Make acid bath (change monthly?)
b. Day before sampling
• Make phenate reagent
• Make Hypochlorous reagent
• acid wash volumetric flasks and pipettes (wash flasks only
before first sampling; afterwards, leave reagents in flasks and
prereact).
• rinse volumetric flasks with DI
• Prereact all volumetric flasks (add fresh reagent each time).
• Prereact 50-ml centrifuge tubes (both standards and sample
tubes). This is a separate step only for the first sample time.
After the first sampling old reagent will be left in the tubes and
used to prereact.
c. Before sampling
• Shake sample tubes
• Dump reagent in proper waste container
• Rinse several times with DI
• Shake dry
• Recap
• Rinse in field with filtered water
d. After sampling
• Place samples in refrigerator while preparing everything
• Shake prereacting volumetric flasks
• Dump reagents in proper waste container
• Rinse several times with DI
• Make stock standard B (10 mol/L in 0.5-L volumetric)
• Calibrate pipetters (if using these)
• Make dilutions. Use the same flask for each standard each
week
• Shake prereacting standard tubes
• Dump reagent
• Rinse several times with DI
• Pour standard in proper tube dedicated to that standard (as clos
e to 20 ml as possible)
68 |
Arctic LTER Streams Protocol
•
•
•
•
•
•
Add reagents from repipetters to standard and sample tubes
Wait an hour or two.
Make sure spec waste is hooked up to right container
Read on spec in order that reagents were added.
Rinse spec w/ di to clean out all reagents
Leave reagents in the flask
E. Nitrate – Lachat Method used at the MBL.
1. This Lachat method (QuikChem (Method 31-107-04-1C ) was used for
Nitrate analysis at the MBL lab. Method 31-107-04-1E is now used at
the RESL
2. Standards for this method are KNO3, opposed to NaKO3 and KNO2,
opposed to KNO2, which is used in the more current method.
Nitrates samples were analyzed on a Lachat Quik-Chem 8000, following QuikChem Method 31-107-04-1-C.
2000. Diamond, David H. Determination of nitrate and/or nitrite in brackish or
seawater by flow injection analysis colorimetry. Quik-Chem Method 31-10704-1-C. Zellweger Ananlytics, Lachat Instrument Division, 6645 West Mill
Road, Milwaukee, WI 53218.
Summary: Samples are passed through a copperized cadmium column to reduce
nitrate to nitrite. The nitrite (both original and reduced nitrate) is determined by
diazotizing with sulfanilamide and coupling with N-(1-napthyl)-ethylenediamine
dihydrochoride to form a highly colored azo dye which is measured
colorimetrically.
Working Range: 5-50µm, MDL 0.12µm
Standard Preparation:
50mM stock standard-in a 1 liter volumetric flask dissolve 0.5055g dried (1hour
at 60°C) potassium nitrate (KNO3) into about 800ml distilled water. Dilute to 1
liter with DI water and invert several times to mix. Can be stored refrigerated for
up to 3 months.
50µm Working Stock Standard-In 1 L volumetric dilute 10ml 50mM stock to
mark with DI water, invert to mix.
II.
69 |
Working Standards (Prepare Daily):
50µm-dilute 250ml 50µm working stock std. with 0ml DI water
25µm-dilute 125ml 50µm working stock std. with 125ml DI water
10µm-dilute 50ml 50µm working stock std. with 200ml DI water
5µm-dilute 25ml 50µm working stock std. with 225ml DI water
0µm-250ml DI water
Arctic LTER Streams Protocol
V.
A.
Appendix B – Locations of Equipment
At Toolik
1. Wet Lab
a. Streams Desk
• Office supplies
• Labels
• Filters, forceps
• Wader patches
• Truck keys and info
• Protocols
• Lab books
b. Streams Shelves - Regularly used field gear
• Nuts and scrubs kits
• Yoy containers and nets
• Wading rods
• Flowtracker
• Batteries
• Label tape
• Ziplock bags
• Caulk gun
c. Streams Closet
• Waders
• Wading boots
• Bug shirts
• Head nets
d. Fluorometer room – dark room
• Ammonium fluorometer
• Ammonium scint vial flats
• OPA reagent and buffer
• Chlorophyll fluorometer
• Borosilicate tubes
• Cooler for Chlorophyll samples
• Acetone
e. Spec room
• Cary spec set up for phosphorus analysis
• Phosphorus working reagent
• Balance
• Vortex
70 |
Arctic LTER Streams Protocol
2. Wet Lab Conex
a. Sample bottles, petri dishes, etc…
b. Wash bottles
c. Amber liter bottles
d. Centrifuge tubes
e. Scint vials
f. Dripper parts and pumps
g. Old field equipment bits
3. Lab 3 – LTER things
a. YOY balance
b. YOY knockout solutions
c. Fishing rods and reels
d. Waders
e. Wading boots
f. Electroshocker
g. Oksrukuyik stream level datalogger
B. At the RESL - 2013
1. RESL Lab
a. Chemicals for reagents, standards
• Ascorbic acid
• Sulfanilamide
• Potassium persulfate
• Potassium nitrate
b. Filter samples
c. Algal comp samples
d. Test tubes
• TDP/PP with polyphenol screw top caps in racks – 15mm and
30mm
• TDN/DOC with septa screw top caps – ash before analysis
• NO3 borosilicate tubes in racks
e. Filters – 47mm and 25mm – some ashed
f. Standard volumetric flasks, other standard glassware.
2. General Use Lab
a. Sodium hydroxide 15N – base cabinet
b. Phosphoric acid – 85% - acid cabinet
c. Sulfuric Acid – 36N – acid cabinet
d. Hydrochloric Acid 12N – acid cabinet
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Arctic LTER Streams Protocol
C. Stored at the MBL
1. Walk in Cooler
a. Water samples from previous summers
• TDP - acidified
• TDN/DOC - acidified
• Anions
• Cations - acidified
• Alkalinity
2. Walk in Freezer
a. Frozen samples from previous summers
• NO3
3. Office
a. Old field notebooks
b. State of the river reports
c. Old reprints
d. Data backup disks
4. Long term storage and memorial circle
a. Old particulate filters
72 |
Arctic LTER Streams Protocol
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