Genetic Sampling Protocol

Collection and storage of non-invasive DNA samples from wild primates

Important aspects to consider
Some general comments:
Collection of fecal material:
1) Storage in 95 % Ethanol
2) Alcohol, followed by silica gel desiccation
3) RNA later
4) 20% DMSO / saturated NaCl2
5) Storage techniques for shed hair:
Suggested reading:

Genetic analyses have become an extremely valuable tool to study multiple aspects of primate evolution. Sources of DNA usually vary between organisms, but usually researchers prefer to obtain DNA samples through non-invasive means, such as fecal material or shed hair. Several studies have shown that storage methods could potentially have a great effect on DNA quality and yield. Here, we describe four storage methods for faecal DNA that has been used by various research groups. We do not intend this list to be all inclusive, but rather focus on methods that allow storage of DNA material in difficult field conditions.

Important aspects to consider

a) Availability of reagents and storage vials in the field
b) Storage conditions in the field
c) Transport of samples from field to laboratory
d) Cost of reagents

We have identified five different protocols. The first three are suited for fecal material, while the forth one works well with small pieces of tissue, if for instance a dead animal is encountered. The last protocol describes the storage and preservation of shed hair.

Some general comments:

Fecal material should be collected as soon as possible after defecation. As a rule of thumb, all samples should be stored as cool as possible (preferably frozen, but (4°C works well, too). Unfortunately, in many field situations this is not possible. A possible solution is a so-called “dry-shipper” (sometimes called a “vapor-shipper”). These are fabricated from durable, lightweight aluminium, containing a hydrophobic absorbent that traps the liquid nitrogen. If properly filled, a dry shipper can cool specimens to -150°C for three weeks, even at high ambient temperatures. Dry-shippers are also approved my most airlines. If refrigerated storage is not possible in the field, the very least that should be done is storing the specimens at a cool and dry place. In areas of high humidity, this could be achieved by placing the samples in a sealable plastic container that contains silica gel to absorb the moisture. Generally, samples should be transferred to a lab as soon as possible.

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Collection of fecal material:

IMPORTANT: Most of the shed cells of the target organism are found at the ‘front end’ and the outside of the fecal material. Hence, it is important to scrape as much as possible from the outside of the material in order to maximize DNA yield. To avoid contamination with human DNA, gloves should be worn at all times when handling samples (although we know that this is sometimes difficult to do in tropical environment). For each sample, a new pair of gloves should be used to minimize the risk of cross-contamination.

1) Storage in 95 % Ethanol

Materials required:

• 8 ml polypropylene tube, lid with o-ring (for instance Sarstedt 60.542.007)
  
• 95% Ethanol or Methanol
  
• Parafilm (www.parafilm.com)
  

Procedure:

• Fill half of polypropylene tube with ethanol or methanol
  
• Place a grape-sized sample in the tube, make sure that it is fully submerged
  
• Seal tube with Parafilm
  
• Label the tube with relevant information
  
• Record relevant information (date, GPS coordinates, sample ID, sex, age, name of location, collector’s name, etc) in a datasheet
  

Advantages:

• Cheap
  
• Quick
  

Disadvantages:

• DNA appears to degrade more quickly compared to other methods
  
• Flammable, might lead to transport problems
  

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2) Alcohol, followed by silica gel desiccation

Materials required:

• 15 ml polypropylene tube with spoon in lid (e.g. Sarstedt)
  
• 8 ml polypropylene tube, lid with o-ring (e.g. Sarstedt 60.542.007)
  
• 95% Ethanol or Methanol
  
• Parafilm (http://www.parafilm.com)
  
• Silica Gel
  
• KimWipes (http://www.kimberly-clark.com/)
  

Procedure:

• Collect fecal sample directly in approximately 15 ml tubes with built-in spoon filled with 5-7 ml alcohol (95%)
  
• Label the tube with relevant information
  
• Record relevant information (date, GPS coordinates, sample ID, sex, age, name of location, collector’s name, etc) in a datasheet
  
• Let sample soak in alcohol for 24-36hrs
  
• After 24-36hrs, transfer 2g fecal sample to two 8 ml tubes
  
• Place small square of KimWipe on sample to separate silica from sample
  
• Add 4g of silica, so silica ends up between sample and lid
  
• Cover silica tubes with Parafilm
  
• Leave remainder of fecal sample in alcohol, add a little extra alcohol to tube for preservation if necessary
  
• Check after 2-3 days whether sample is dry.
  

If yes: Monitor silica gel every two-three weeks for change in color, replace silica gel ONLY if necessary
If not dry: replace silica gel, check again after two to three days.

Advantages:

• DNA yield and quality is higher compared to method 1
  
• Samples are dry and non-flammable
  

Disadvantages:

• Repeated handling increases contamination risk
  
• More expensive than method 1
  
• Several different tubes and containers are required
  

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3) RNA later

RNA later (manufactured by several companies, e.g. Ambion, Sigma-Aldrich, Qiagen and others!) is a specialized storage solution that was originally designed to preserve RNA in samples collected in the field. It appears to preserve DNA very well, and seems to have a superior performance compared to methods 1 and 2. Unfortunately, it is not cheap.

Materials required:

• 8 ml polypropylene tube, lid with o-ring (e.g. Sarstedt 60.542.007)
  
• RNA later solution (Ambion # 7020, Sigma-Aldrich #R0901)
  

Procedure:

• Add 3ml of RNA later to the tube
  
• Place a grape-sized sample in the tube, make sure that it is fully submerged
  
• Seal tube with Parafilm
  
• Label the tube with relevant information
  
• Record relevant information (date, GPS coordinates, sample ID, sex, age, name of location, collector’s name, etc) in a datasheet
  

Advantages:

• DNA yield and quality higher than for methods 1 and 2
  
• Simple
  
• Small contamination risk
  
• Non flammable
  

Disadvantages:

• Expensive (~US$ 2.50 / sample)
  

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4) 20% DMSO / saturated NaCl2

This method works very well to preserve small quantities of tissue at room temperature. It is basically like pickling meat. The DMSO helps to rupture cell structures, so the salt can penetrate the cells much quicker to denature proteins.

Materials required:

• 8 ml polypropylene tube, lid with o-ring (e.g. Sarstedt 60.542.007)
  
• 20% (v/v) di-methyl sulphoxide (DMSO) / saturated (5 M) NaCl2 solution (needs to made before sampling)
  
• Parafilm
  

Procedure:

• Add 4ml of DMSO/sat. NaCl2 to tube
  
• Place a grape-sized sample in the tube, make sure that it is fully submerged
  
• Seal tube with Parafilm
  
• Can be stored at room temperature for several months
  
• Label the tube with relevant information
  
• Record relevant information (date, GPS coordinates, sample ID, sex, age, name of location, collector’s name, etc) in a datasheet
  

Advantages:

• Cheap
  
• Non-flammable
  

Disadvantages:

• Needs to be prepared ahead of field work
  
• Not suitable for fecal material
  

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5) Storage techniques for shed hair:

Materials required:

• Gloves (one pair per sample)
  
• Tweezers
  
• Lighter
  
• Paper envelope
  
• Silica gel
  

Procedure:

• Wear gloves
  
• Collect the hairs with tweezers that are sterilized with a flame between each use.
  
• Place hairs into a clean paper envelope
  
• Place the envelopes in a plastic bag (Ziploc) together with silica gel (if possible)
  
• Record the following information: date, GPS coordinates, sample ID, nest age, height of nest from ground, collector’s initials, and reference to any faecal samples collected nearby.
  

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We thank Todd Dissotell for providing information regarding the RNA later storage method.

Suggested reading:

Goossens B, Anthony N, Jeffery K, Johnson-Bawe M and Bruford MW (2003). Collection, storage and analysis of non-invasive genetic material in primate biology. In Setchell JM, Curtis DC, ed. Field and Laboratory Methods in Primatology, pp. 295-308. Cambridge University Press, Cambridge.

Jeffery KJ, Abernethy KA, Tutin CEG, Bruford MW (2006) Biological and environmental degradation of Gorilla hair and microsatellite amplification success in hair extracted DNA. Biological Journal of the Linnean Society, in press.

Murphy MA, Waits LP, Kendall KC (2000) Quantitative evaluation of fecal drying methods for brown bear DNA analysis. Wildlife Society Bulletin, 28, 951-957.

Murphy MA, Waits LP, Kendall KC, Wasser SK, Higbee JA, Bogden R (2002) An evaluation of long-term preservation methods for brown bear (Ursus arctos) faecal DNA samples. Conservation Genetics, 3, 435-440.

Nsubuga AM, Robbins MM, Roeder AD, Morin PA, Boesch C, Vigilant L (2004) Factors affecting the amount of genomic DNA extracted from ape faeces and the identification of an improved sample storage method. Molecular Ecology, 13, 2089-2094.

Roeder AD, Archer FI, Poinar HN, Morin PA (2004) A novel method for collection and preservation of faeces for genetic studies. Molecular Ecology Notes, 4, 761-764.

Valderrama X, Karesh WB, Wildman DE, Melnick DJ (1999) Non-invasive methods for collecting fresh hair tissue. Molecular Ecology, 8, 1749-1752.