Field preservation of marine invertebrate tissue for DNA analyses

Field preservation of marine invertebrate tissue for DNA analyses
Molecular Marine Biology and Biotechnology (1998) 7(2), 145–152
Field preservation of marine invertebrate tissue
for DNA analyses
Mike N Dawson*, Kevin A. Raskoff, and
David K. Jacobs
Department of Biology, UCLA, P.O. Box 951606,
Los Angeles, California 90095-1606, U.S.A.
Successful preservation of tissue samples is a prerequisite for long field studies in remote areas.
However, there is little published information concerning field preservation of marine invertebrate
tissues for DNA analyses. This omission is significant because marine biodiversity is centered in the
Indo-Pacific, where immediate DNA analysis is often impossible. Consequently, we used an assay
based on polymerase chain reaction (PCR) to examine the effect of five storage solutions and three
temperature regimens on the degradation of DNA
from four common classes of marine invertebrates
(Anthozoa, Gastropoda, Polychaeta, and Scyphozoa). Control samples were cryopreserved. Storage
solution and the type of tissue preserved were the
best predictors of preservation success. The length
of time in storage and the storage temperature also
affected the preservation of DNA. A field test demonstrates that a solution of dimethylsulfoxide and
sodium chloride (DMSO-NaCl) preserves a wide
range of tissues for DNA analyses and is very
simple to use in remote field locations.
Plans to investigate scyphozoan phylogenies in the
western equatorial Pacific Ocean required the preservation of tissue samples for long periods of time
in a hot and humid climate. A survey of the literature revealed that, although many studies have
identified methods suitable for preserving plant
and animal tissues (Table 1), there were no recom-
* Correspondence should be sent to this author.
r 1998 Springer-Verlag New York Inc.
mendations on how best to preserve marine invertebrate tissues for DNA analyses. This as a significant omission for several reasons. First, DNA analyses are invaluable in studies of the evolution,
systematics, and population genetics of marine invertebrates (e.g., see McMillan et al., 1991; Avise,
1994, p. 154; Burton and Lee, 1994; Palumbi, 1994;
France et al., 1996). Second, marine invertebrates
are becoming increasingly important to the pharmaceutical industry (Colin and Arneson, 1995). Finally, the marine environment harbors the greatest
diversity of invertebrates (Brusca and Brusca, 1990,
p. 5), and this diversity is highest in the IndoPacific (Colin and Arneson, 1995), where immediate analysis, or cryopreservation, of DNA is often
DNA is particularly susceptible to degradation
by hydrolytic and oxidative endogenous nucleases
(Dessauer et al., 1995), which, if not countered,
break down highly informative long strands of
DNA into small fragments of greatly reduced use
for many analyses (Seutin et al., 1991). Enzyme activity, and consequently DNA degradation, may be
limited by adjusting the ambient pH, salt concentration, or temperature (Dixon and Webb, 1979).
Samples may be successfully preserved by a number of chemical or physical treatments (Table 1).
Cryopreservation is the preferred method of DNA
protection (Chase and Hills, 1991; Seutin et al.,
1991; Rogstad, 1992; Post et al., 1993; Reiss et al.,
1995), and may be accomplished by freezing
samples over dry ice (−78°C) or in liquid nitrogen
(−196°C). However, deep freezing is not always feasible. Both dry ice and liquid nitrogen are difficult
to use in the field because they require careful handling and special equipment, and furthermore,
strict regulations limit their transport by air (Liston
and Rieseberg, 1990; Chase and Hills, 1991; Seutin
et al., 1991; Dessauer et al., 1995).
This study was undertaken to identify an alternative to cryopreservation, suitable for the longterm storage of marine invertebrate tissues for DNA
analyses, and appropriate for use at remote field
sites. After reviewing the published literature, we
M.N. Dawson, K.A. Raskoff, and D.K. Jacobs
Table 1.
A synopsis of DNA-preservation methods.*
Plant leaf
Fly tissue
Wasp tissue
Beetle tissue
Ant tissue
Fish tissue
Bird tissue
Bird blood
Human blood
Storage treatment
x f, ug
* In the body of the table, a check mark (u) indicates successful preservation of DNA; x, degradation of DNA; *, conflicting results depending on
precise protocol; blank cell, no information available; superscript numbers, % ethanol (EtOH); DNAB, DNA isolation buffer; superscript letters,
other chemical treatments.
Chemical treatments: aformalin, glutaraldehyde, EDTA, trichloroacetic acid, clorox, 25% NaCl, methanol/chloroform/proprionic acid, perfix;
propanol, methanol, Carnoy fixative, formal saline; cethyl acetate, Carnoy fixative; dchelating ion-exchange resin; ephenol; fformalin; gethylene
Reference numbers: 1Pyle and Adams (1989), Chase and Hills (1991), Rogstad (1992), Nickrent (1994), Flournoy et al. (1996); 2Post et al. (1993);
Dillon et al. (1996); 4Reiss et al. (1995); 5Altschmied et al. (1997); 6Proebstel et al. (1993), Asahida et al. (1996), Altschmied et al. (1997); 7Seutin
et al. (1991); 8Albariño and Romankowski (1989); 9Holzmann and Pawlowski (1996).
chose to investigate the effects of five buffer solutions (70% ethanol, ‘‘Queen’s’’ lysis buffer—see Experimental Procedures, DMSO-NaCl solution,
hexadecyl trimethyl ammonium bromide [CTAB]–
NaCl solution, and a urea extraction buffer) and
storage at three temperatures (frozen, refrigerated,
and ambient) on the long-term preservation of tissue from four marine invertebrate species. Desiccation was not included in this investigation because
the high water content of many marine invertebrates (e.g., Scyphozoa, Polychaeta) would not be
compatible with the requirement that samples be
dried within 12 h to prevent degradation of DNA
(Chase and Hills, 1991). The species were chosen to
represent four classes of common marine invertebrates: gastropod mollusks (Astraea undosa), polychaete worms (Phragmatopoma californica), and
two cnidarians Anthopleura xanthogrammica
(anemone) and Aurelia sp. (scyphozoan jellyfish).
Tissue from all species was stored in each solution
and placed in all temperatures for up to 28 months
prior to analyses. Control samples were cryopreserved in the absence of storage solutions. From the
results of the experimental study, we decided to
undertake a field test of DMSO-NaCl.
The preservative used and the type of tissue preserved were the two principal factors influencing
the degradation of sample quality. Lesser effects
were attributable to the temperature and duration
of storage (Figure 1A).
Visible physical degradation was negligible in
all samples stored at −80°C. The physical appearance of tissues degraded in almost all other treatments (Figure 1A). Of these treatments, physical
structure was preserved best by DMSO-NaCl and
70% ethanol; almost without exception structure
could be easily identified after 28 months of storage
in these solutions. In contrast, storage in urea resulted in complete dissolution of most samples
during 28 months, the principal exceptions were
those samples that were frozen. The Queen’s and
CTAB-NaCl methods were intermediate in performance—neither satisfactorily preserved the physical structure of samples. Typically, and regardless
of preservative, degrading Phragmatopoma appeared as skeleton-like structures that would break
up if disturbed. Degrading tissues of Aurelia became a slurry in the bottom of all tubes. Degradation of Astraea and Anthopleura, when not dissolved by urea, was commonly apparent as a loss of
opacity of the tissues.
The physical condition of a sample was a poor
indicator of the quality of DNA in that sample. High
molecular weight DNA (∼20 kb) was extracted from
most samples stored for up to 28 months in all solutions or at −80°C, although considerable degradation of DNA also occurred in some of these samples
Field preservation of marine invertebrates for DNA analyses 147
Figure 1A. Effect of storage treatments on the preservation of tissue samples for DNA analyses. Each column represents
a combination of one storage solution and one storage temperature (−13°C, 6°C, 25°C), excepting ‘‘−80,’’ which indicates
storage at −80°C alone. Within these columns, each cell indicates the effect of that treatment on the preservation of one
of four tissue types, stored for 1, 6, or 28 months, assessed by three criteria: (1) preservation of the visual appearance of
tissues, assessed on a 4-point scale from black (well preserved) through to white (poorly preserved); (2) the latest stage
(months) at which high molecular weight DNA was successfully extracted from the sample; (3) the latest stage at which
DNA extracts were successfully amplified using PCR. Amplifications using DNA from Phragmatopoma stored for 1
month were all unsuccessful. This is attributed to degradation of the DNA after extraction because fragments were
successfully amplified from Phragmatopoma samples stored for 6 months. Asterisks indicate significant degradation of
DNA although some high molecular weight DNA is still present.
Figure 1B. Examples of successful and unsuccessful extractions of high molecular weight DNA. **Successful: lane 1
(Anthopleura, stored in CTAB-NaCl at 6°C for 28 mo) and lane 2 (Anthopleura, DMSO-NaCl, 6°C, 28 mo). *Successful
with degradation: lane 3 (Anthopleura, 70% ethanol, 25°C, 28 mo) and lane 4 (Astraea, Queen’s, 6°C, 28 mo). Unsuccessful: lane 5 (Astraea, Queen’s, 25°C, 28 mo). M indicates 100-bp ladder.
Figure 1C. Examples of successful (**) and unsuccessful amplification of the ITS-1 region by PCR. All products were
amplified from Astraea stored at −13°C for 28 months; ‘‘—’’ denotes a negative control.
(Figures 1A, 1B). The failure of almost all treatments to prevent considerable degradation of DNA
in samples of Aurelia is conspicuous.
The success of PCR amplification primarily reflected effects attributable to tissue type (Figures
1A, 1C). With few exceptions, DNA extracted from
Astraea and Anthopleura was successfully ampli-
fied at each stage of the 28-month investigation.
Amplifications of Aurelia DNA showed far greater
variability in success and little pattern with regard
to preservation method. Attempts to amplify ITS-1
from Phragmatopoma DNA, however, were more
consistently unsuccessful. DNAs extracted from
Phragmatopoma after 1 month and 28 months in
M.N. Dawson, K.A. Raskoff, and D.K. Jacobs
storage all failed to amplify; DNA extracted after 6
months in storage did yield amplification product
but not in a predictable manner from any particular
treatment. Of those samples stored at −80°C, only
Phragmatopoma DNA failed to amplify after 28
months in storage.
These results show that the principal factors affecting sample preservation were tissue type and
storage buffer. If tissue was not susceptible to degradation (Astraea and Anthopleura), then the storage solution had little effect on the preservation of
DNA. Conversely, if samples were susceptible to
degradation (Aurelia and Phragmatopoma), then
the storage solution used had considerable influence on the success of preservation (Figure 1A). In
these cases, preservation using CTAB-NaCl,
Queen’s, or urea was noticeably less successful
than treatments using DMSO-NaCl or 70% ethanol.
Two other factors, duration and temperature of
storage, also influenced the success of sample preservation. First, visual inspection of samples stored
for 1, 6, and 28 months suggests that the greater the
duration of storage prior to analysis, the greater the
degradation of the sample. This pattern was most
obvious from the visual assessments of physical
quality, but was also reflected in increasing fragmentation of high molecular weight DNA with
time. Second, in the few cases in which preservation success varies according to the storage temperature, it seems that greater degradation occurs at
the higher storage temperature (e.g., see Aurelia),
although this effect is neither consistent nor pervasive (and is arguably reversed in Phragmatopoma).
Notably, temperature affected the appearance of
some storage solutions. Refrigeration and freezing
caused both CTAB-NaCl and urea solutions to separate into two layers (one white and one clear),
while Queen’s solution froze when kept at −13°C.
The physical appearances of DMSO-NaCl and 70%
ethanol were not affected by storage temperature.
Samples of marine invertebrates and fish collected in Palau and stored under refrigeration in
DMSO-NaCl for between 6 and 18 months were in
good physical condition. Most provided high molecular weight DNA, and all but one were amplified
by the PCR using either cytochrome c oxidase subunit I or ‘‘D-Loop’’ primers (Figure 2).
Many factors affect the preservation of DNA, including the type of tissue, the chemical and physical environment in which that tissue is stored, and
the duration of storage. However, the interactions
of these factors and their resultant effects on DNA
preservation are difficult to predict a priori. Consequently, one might wish to test several alternative
methods prior to field collections, or to use more
than one preservation method once in the field
(Chase and Hills, 1991; Rogstad, 1992). However, in
the absence of such contingency plans, this research suggests that storage in DMSO-NaCl is the
method most likely to result in successful preservation of tissue samples. Added precautions include storing the preserved samples at reduced
temperature and performing DNA extractions at the
earliest opportunity.
The most striking result of this study is the effect
of different types of tissue on the success of preservation. These differences may be due to several
factors. For example, samples of the two species
that were most successfully preserved, Astraea and
Anthopleura, were taken from muscular tissue,
which is physically more robust than either the soft
body of Phragmatopoma or the very delicate gonadal and gastric tissues of Aurelia. Second, a
greater concentration of catalytic enzymes was
most likely present in samples of Phragmatopoma
and Aurelia than in samples of Astraea and Anthopleura as the former samples both included gastric
material. Finally, some constituent of Phragmatopoma was responsible for making the successfully
preserved and extracted high molecular weight
DNA unavailable for amplification. Tissue type has
also been found to affect the success of preservation
and DNA analyses of samples from plants (Pyle and
Adams, 1989; Chase & Hills, 1991; Rogstad, 1992)
and birds (Seutin et al., 1991), and Altschmied et
al. (1997) reported the abdomen of some ants must
be discarded prior to preservation else the formic
acid therein will depurinate the DNA. Other chemical interactions affecting the recovery of high molecular weight DNA have been noted by Flournoy
et al. (1996).
In addition to tissue type, the choice of storage
solution exerted an important influence on the rate
of degradation of samples. In accord with Seutin et
al. (1991), we found that DMSO-NaCl was the best
solution in which to store tissues. This was the
only solution that preserved recognizable tissue
and amplifiable high molecular weight DNA from
Aurelia. DMSO-NaCl may protect DNA in several
ways. DMSO is a cryoprotectant (Dessauer et al.,
1995) and thus prevents freeze-thaw damage of
samples, although damage incurred in this way
may not be a significant concern (Seutin et al.,
Field preservation of marine invertebrates for DNA analyses 149
Figure 2. Results from a field test of the DMSO-NaCl preservation method. (A) DNA extracts from tissues of 9 taxa stored
in DMSO-NaCl at 5°C for 6–18 months. (B) PCR products amplified from these DNA extracts.
1991). Also, by perturbing the structure of membrane-bound proteins, DMSO enhances the absorption into cells of materials, such as EDTA and NaCl,
that inhibit nucleases (Seutin et al., 1991). The utility of high-salt solutions in preserving DNA was
also demonstrated by the CTAB-NaCl treatment, although this method was less successful than preservation in 70% ethanol. Storage in ethanol dehydrates the sample and results in the denaturation
and precipitation of proteins, including catabolic
enzymes (Flournoy et al., 1996).
This study did produce unexpected results.
First, the urea-based solution was suitable for longterm storage of DNA despite the assertion of Seutin
et al. (1989) that, within 6 months, urea transforms
into ammonia resulting in an elevated pH in which
DNA is denatured. Asahida et al. (1996) have also
found a urea-based preservative suitable for long-
M.N. Dawson, K.A. Raskoff, and D.K. Jacobs
term storage of tissues for DNA analyses. Second,
preservation of DNA in lysis solutions (urea and
Queen’s) was arguably unaffected, or even improved, by lower storage temperatures although
such buffers were designed for use at ambient temperature (Seutin et al., 1991). Clearly, any decline
in the efficacy of these preservatives is, at least,
offset by benefits of storage at lower temperatures.
Third, these ‘‘lysis’’ solutions preserved DNA effectively within the stored tissue even though their
intention is to lyse cells and release DNA into the
surrounding solution (Seutin et al., 1991); cell lysis
is not necessarily synonymous with movement of
DNA from that cell into solution, particularly if the
lysed cell is in the midst of a largely intact tissue
Compared with tissue type and storage solution,
the duration and temperature of storage had minor
effects on the degradation of samples over the time
course of this study. However, these minor effects
agree with previous investigations that reported the
quantity and quality of DNA recovered from
samples progressively declines as the duration of
storage increases (Post et al., 1993; Reiss et al.,
1995), and that DNA degrades less rapidly in colder
environments (Post et al., 1993; Poinar et al., 1996).
Preservation of tissue may be facilitated by
finely dicing tissue to increase permeation of the
storage solution into the sample (Seutin et al., 1991;
Reiss et al., 1995). Dessauer et al. (1995) suggest
that tissue should be minced into pieces no larger
than 1 mm.3 However, none of our samples that
yielded high molecular weight DNA contained
pieces less than 1 mm3, with the exception of gastric filaments and gonads of Aurelia. Further, the
preservation of intact physical structure is often desirable because samples may then be used for morphologic or parasitologic analyses as well as DNA
analyses (Post et al., 1993; Reiss et al., 1995). Both
physical structure and DNA were preserved well by
DMSO-NaCl and 70% ethanol.
Although reproducing results has been a problem in previous studies (see Post et al., 1993), we
experienced few discrepancies between independent tests, run concurrently by two of the authors.
Consequently, the general patterns apparent in this
study are expected to be robust to further investigation. The discrepancies that did occur were predominantly in species and storage groups that exhibited degradation in at least several treatments.
In contrast, there were no contradictory results for
any sample stored in DMSO-NaCl. This implies
that, of the treatments investigated, storage in
DMSO-NaCl is least sensitive to small inconsistencies (e.g., precise source and volume of tissue) that
may occur during the preservation of samples.
Field testing of the DMSO-NaCl method supported the experimental results and also showed
that this method is appropriate for a wide range of
organisms and is extremely simple to use in the
field. The DMSO-NaCl solution may be made and
aliquoted into microcentrifuge tubes prior to departure. These tubes may be kept at any moderate temperature, before and after they are used to preserve
samples, although chilling is preferable. The few
items that need to be taken on each field excursion
are the appropriate number of prepared microcentrifuge tubes, a small dissection kit with which to
collect the sample, and materials such as sterile
water, ethanol, and a cigarette lighter with which to
clean samples and sterilize equipment. This small
volume and mass of equipment is appropriate for
lengthy collecting trips to remote sites.
Experimental Procedures
Collections were made on the coast of California
between June 19 and 26, 1995. Tissues from Aurelia sp. were extracted at the field site and placed
into storage solutions in temporary temperature
conditions for the duration of sampling and transport to UCLA (<10 h). During sampling and transportation, deep-frozen, frozen, and refrigerated
samples were kept on ice, while room-temperature
samples were kept at ambient temperature. Astraea
undosa, Phragmatopoma californica, and Anthopleura xanthogrammica were transported live to
UCLA where they were kept in a seawater aquarium until tissues were dissected out.
Tissue samples, approximately 0.2 cm3, were
taken from Phragmatopoma californica (a single
whole worm), Astraea undosa (foot muscle), Anthopleura xanthogrammica (body wall), and Aurelia sp. (gonad and stomach). Each sample was
washed with 0.22 µm filtered seawater and then
deposited into 500 µl of each of five storage solutions. The saturated NaCl-CTAB solution described
by Rogstad (1992) was modified to consist of 0.1 M
Tris (pH 8.0), 0.02 M EDTA (pH 8.0), 0.02% (wt/
vol) CTAB and saturated with NaCl. This solution
was autoclaved and cooled prior to the addition of
b-mercaptoethanol to 0.002%. DMSO-NaCl solution is 20% DMSO, 0.25 M disodium-EDTA, and
NaCl to saturation, pH 7.5 (Seutin et al., 1991); the
pH of this solution must be above 8 for the EDTA to
dissolve, and warming promotes dissolution of
Field preservation of marine invertebrates for DNA analyses 151
NaCl. Queen’s lysis buffer contains 0.01 M Tris,
0.01 M NaCl, 0.01 M disodium-EDTA, and 1.0%
n-lauroylsarcosine, pH 8.0 (Seutin et al., 1991).
Both DMSO-NaCl and Queen’s solutions were sterilized by autoclaving prior to use. Urea buffer (168
g of urea, 25 ml of 5 M NaCl, 20 ml of 1 M Tris, 16
ml of 0.5 M EDTA, 40 ml of 10% sodium dodecyl
sulfate (SDS), plus 170 ml of distilled water, pH
8.0) was filter-sterilized under vacuum through a
0.2-µm millipore filter; 100% ethanol was diluted
to 70% using 4.0-µm millipore filtered seawater
and then autoclaved.
Replicate samples of each of the 20 possible
combinations of species tissue and storage solution
were kept under three temperature regimens (mean
± 2.5°C): frozen (−13°C), refrigerated (6°C), and ambient (25°C). In addition to these experimental conditions, control samples of each tissue, with no
added storage solution, were cryopreserved at
−80°C. All samples were stored in opaque containers to prevent damage of samples by UV irradiation
(Dessauer et al., 1995).
After 1 and 6 months in storage, two of the authors (K.A.R. and M.ND.) independently assessed
the state of tissues stored in each of these conditions. The analyses at 28 months were performed
solely by M.ND. First, the physical condition of
tissues was visually assessed according to a 4-point
scale, by comparing the appearance of stored tissue
relative to the appearance of fresh material. Second, each treatment was scored for the presence or
absence of high molecular weight DNA. Amplification of a PCR product from the extracted DNA was
the third indicator of a successful preservation
In general, tissue pelleted out of the storage solution by centrifugation for 5 min at 14,000 rpm
was used for the DNA extraction. The supernatant
was pipetted off (see below*), and the pelleted tissue was resuspended in 600 µl CTAB (0.1 M Tris
[pH 8.0], 0.02 M EDTA [ph 8.0], 0.02% [wt/vol]
CTAB, 0.8 M NaCl, 0.002% b-mercaptoethanol)
with 6 µl proteinase K (20 mg ml−1) and digested at
42°C for 16 h. This proteinase K digestion was
omitted for samples from Aurelia. DNA is liberated
from these tissues by grinding in a dounce. Digested or ground samples were centrifuged for 5
min at 14,000 rpm before 300 µl of the supernatant
was taken for DNA extraction. If samples were
stored in urea or Queen’s, however, only 150 µl of
this supernatant was used in the DNA extraction;
the other 150 µl was pipetted from the original preservative (see above*). When no tissue was pre-
served by urea or Queen’s, 300 µl of the original
preservative was used in the DNA extraction. The
DNA extraction consisted of a single extraction
with chloroform followed by repeated extractions
with phenol:chloroform:isoamyl alcohol (25:24:1)
until the interface between aqueous and organic
phases was clear. A single chloroform:isoamyl alcohol (24:1) extraction was then completed before
precipitating the DNA at −20°C for 1 h with 3 M
sodium acetate and 100% ethanol. Purified DNA
was resuspended in 30 µl of sterile water after the
precipitate had been centrifuged for 28 min at
13,000 rpm, 6°C, and the resulting pellet was
washed in 75% ethanol and dried at 25°C. The
quality of DNA extract was determined by electrophoresing an 8-µm aliquot of each DNA sample
across a 1.4% agarose minigel (1% TBE) containing
ethidium bromide and scoring for either presence
or absence of high molecular weight DNA. The
ITS-1 region was amplified using the primers
(Vogler and DeSalle, 1994). Approximately 1 ng of
sample was added to each 25 µl total volume for
PCR, set up according to the guidelines issued with
Taq polymerase (Perkin Elmer). A ‘‘hotstart’’ was
included before entering a 28-cycle PCR. Each
cycle on the MJ Research MiniCycler comprised 45
s at 94°C, 45 s at 50°C, and 60 s at 72°C. PCR product was visualized as above, and scored for either
presence or absence of a single strong band within
the size range of previously sequenced ITS-1 regions (Vogler and DeSalle, 1994).
Based on preliminary (1-month) results, tissues
from 10 additional taxa were sampled and stored
for 6 to 18 months in DMSO-NaCl (∼5°C) during
field trips to the Republic of Palau, Micronesia, in
1996/1997 (6 mo, mussel, nereid polychaete; 9 mo,
gammarid amphipod, goby, cardinalfish; 10 mo,
anemone, Aurelia, shipworm, scaleworm; 18 mo,
mastigiid jellyfish). All steps of the subsequent
DNA extraction, PCR, and visualization of products
were completed as above. At this time, other primers were used to amplify the mitochondrial controlregion
Lee et al., 1995), or cytochrome c oxidase subunit I
M.N. Dawson, K.A. Raskoff, and D.K. Jacobs
Folmer et al., 1994) from these samples.
Funding for this research was supplied by a grant
from the Committee on Research of the Academic
Senate of the Los Angeles Division of the University of California. Thanks to Charlie Wray for supplying the ITS-1 primers, to Ted Groscholz for use
of laboratory facilities at Bodega Bay Marine Laboratory, and to the Coral Reef Research Foundation
for providing boats and laboratory facilities in
Palau. Two anonymous reviewers provided comments that resulted in the improvement of this
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