Difference between revisions of "Tissue Sample Collection"

Revision as of 17:37, 13 May 2019

Statement of Purpose

These links and documents contain information about best practices for tissue sample collection

Introduction

Contributors

Major editor: Breda Zimkus. Original content generated during The American Society of Ichthyologists and Herpetologists (ASIH) Annual Joint Meeting - 2016, during an iDigBio sponsored workshop by the following individuals participating in the "Field to Database" Group of the aforementioned workshop: Breda Zimkus, Cesar Aguilar, Ben Frable, Meredith Mahoney, Zachary Randall, and David Wernecke.

Preparation

• Researchers collecting samples need to consider the location of initial preservation and if possible carry out a feasibility study to ensure that the selected preservation method(s) will work given any logistical constraints.
• Sterilization of all dissection equipment (including surface if not using disposable for surface)
• Weighing paper, scraps of thermal paper (left over from printing tags), parafilm, paper towels may be used as disposable surface covers, though none are sterile. Can use on top of cutting board.
• Gloves
• Field catalog to write additional notes (see Preparation)

Vial Selection

One area of immense variability is with the number of vial types used in natural history collections (Zimkus and Ford 2014).

• Samples stored using LN₂ must have vials that are rated for use with cryogenic temperatures, preferably made of polypropylene with screw-top caps. Glass vials are problematic when frozen because of their fragility when handling and vulnerability to cracking when stressed, and their use with LN₂ is unacceptable because leakage into the vials can lead to the vessels exploding.
• Pop-off lids are not recommended because this vial type can easily open on its own.
• Vials can have threads located internally or externally on the vial opening; both vial types exhibit advantages and disadvantages.

- Externally-threaded closures promoteb more sterile conditions because internal threading can allow contaminants to enter if the cap is placed on an unclean surface when removed, but these vials can be susceptible to cracks and loss of air-tightness, which can lead to sample dessication or oxidation (Corthals and DeSalle 2005, Corthals 2006).

-Internally-threaded vials might allow increased storage capacity, depending on the vial selected, but users also suggest that material can be trapped within the threads of this vial type (Johnson 1999).

• Some manufacturers suggest that vial caps that incorporate a silicone gasket or O-ring (internally or externally threaded) are ideal for vapor-phase LN2 freezing because the seal is enhanced, but care must be taken if the vial cap is over-tightened because the gasket can become distended. In addition, the presence of gaskets or O-ring might improve the initial performance of seals but can be problematic when used with some alcohols (e.g., ethanol) because some gasket material, such as silicone, is vapor permeable.
• When working with liquid-phase nitrogen cold storage, extra caution must be taken because the accidental entrapment of liquefied nitrogen inside the vial leads to pressure build up and, upon removal, rapid vaporization of the liquid can result in leakage or even explosion.

- As a precaution, vial manufacturers recommend that samples not be immersed in LN₂ unless they have secondary containment. Heat-sealing vials into flexible polyethylene tubing is recommended for safe storage in the liquid-phase environment.

Labeling and organizing vials

• Pre-label vials

-Many include specimen tags inside vials, but paper inside vials not ideal because of possible contamination

• Use alcohol-proof pen
• Use scribe to etch identifying information as back-up
• Organizing vials numerically in boxes is preferable to storage in bags in case writing comes off vials
• Tissue samples should be taken as soon after euthanization as possible

Vial Considerations

• Internally threaded fit more/box (100 vials/box versus 81 vials/box) but if overfilled can be difficult to open
• Silicone o-ring is vapor permeable (Stuart and Hogue, 2012)

Tissue Collection Methods

Some general recommendations include:

• Collect sample as soon after euthanasia as possible

-Pre-labeling vials may increase preparation speed

• Fill two sets of tissue vials in the field: one for current project and one for institutional collection (rather than subsampling one vial later).
• Label vial to indicate unique number and tissue type

- Only place one tissue type in each vial

- Epigenetic studies require same tissue type for all samples in study (i.e., all muscle)

• Cut/slice up tissue sample, small size allows permeation by preservative (Note: read specific recommendations for preservative provided by manufacturer.)
• Fresh preservative may be needed if in field for extended period of time
• Don’t over-fill vial with tissue, or tissue and preservative (expansion, can interfere with vial threads, etc.)
• NGS methods need reasonable sample size to yield lots of DNA, ~500 ng (my notes say 500 ng, but is that the tissue or the desired amount of DNA? I’m guessing DNA)

Tissue Preservation Methods

The methods currently used to preserve genetic material during initial sampling vary widely owing to the sampling location, tissue type, and intended research use (Dessauer and Hafner 1984, Prendini et al. 2002, Bus´ and Allen 2014). A comprehensive treatise of methods to preserve genetic samples is discussed in Nagy (2010). In a survey of genetic resource collections associated with natural history museums, samples were found to be initially preserved in a variety of different ways (Zimkus and Ford 2014).

Freezing and Flash-freezing

• Freezing samples without the use of a preservation agent either by placing them into a laboratory freezer (≈-20 °C) or by

using ice within an insulated container is generally not recommended because temperatures are not low enough to prevent enzymatic activity nor the formation of intracellular ice crystals (Stoycheva et al. 2007; Nagy 2010).

• Those collecting samples may have access to mechanical freezers that can maintain samples at ultracold or cryogenic temperatures (-80 °C to -150 °C), but this generally excludes samples that are collected in the field.
• Dry ice (frozen carbon dioxide; -78.5 °C) stored within insulated containers may allow the collection or transport of samples, but a sublimation rate of approximately 10% or 2–5 kg every 24 hours limits this method to relatively short trips. In addition, since temperatures near the upper end of the ultra-low temperature range, some DNA degradation may occur as a result of weak enzymatic activity (Nagy 2010).
• Cryogenic storage dewars, a specialized type of vacuum flask used to store cryogenic fluids, can be used to flash-freeze samples in LN₂.

- Access to LN₂ increases field sample collection options, including the preservation of tissues useful for cell culture, RNA, and gametes for multiple weeks.

- The use of LN2 requires additional precautions as it can cause frostbite, cold burns, and asphyxiation by displacing the oxygen of the surrounding area.

-Sample vials can shatter when removed from storage because LN₂ can enter the vials and rapidly expand upon warming, creating a hazard from both flying debris and exposure to the contents. Secondary containment (e.g., polyethylene tubing, tin foil) and protective eyewear is, therefore, recommended.

- Cross-contamination has been reported for samples immersed directly in LN₂, so researchers should consider vial type, secondary containment options, and use of vapor-phase storage when considering flash-freezing methods (Clark 1999).

-Freezing without the inclusion of a preservative was once thought to maximize future research potential, but data now suggests that buffered samples or those stored in a cryoprotectant, such as DMSO or glycerol, may perform better after thawing and refreezing (Nagy 2010)

-Cryoprotectants partly protect against degradation occurring during temperature changes, such as freeze-thaw cycles. If viable cells are desired (e.g., cell culture, gametes), slow freezing and using a cryoprotectant is required to prevent the formation of ice crystals that lead to fatal cell lysis.

• A dry shipper is an insulated cryogenic flask/container that contains LN₂ absorbed into a porous lining.

- Dry shippers are not considered a dangerous product and hence can be used to ship samples by plane if the liquid is fully absorbed and excess poured off.

- There are wide variations in dry shippers with regard to size and temperature (static) hold times.

- Size ranges include those with space for a dozen vials to others that can accommodate thousands of vials.

- Temperature hold times can vary from a few days to multiple weeks, and variations also exist in how well they hold temperature under different environmental and handling conditions.

Ethanol

• Using ethanol as a preservative has numerous advantages for researchers whose primary goal is to preserve DNA.
• Ethanol is easy to use and able to preserve DNA even in areas with elevated ambient temperatures for long periods of time.
• The transport of non-infectious ethanol-preserved specimens has been allowed since 2011 via International Air Transport Association (IATA) Special Provision A180, making it possible to transport specimens preserved in ethanol. See Shipping and Handling of Dangerous Goods for more information about proper packing and labeling.
• Ethanol concentration can greatly affect the resulting quality of the samples with 95–96% (190 proof) recommended as optimal.

- Concentrations above 96% (including absolute ethanol) are not recommended as they likely contain traces of drying agents (e.g., benzene) that can affect DNA preservation (Ito 1992).

- Concentrations of 65–75% (commonly used to preserve whole animals for morphology) are not recommended; Seutin et al. (1991) were unable to recover DNA from bird brain and muscle samples kept in 70% ethanol for six weeks at room temperature, while liver samples yielded significantly degraded DNA.

- Nagy (2010) suggested that tissues be cut into small pieces to increase the surface area, using at least 5:1 volumes ethanol, while others suggest higher ratios (Martin 1977).

- Although the initial concentration and ratio of ethanol to sample is important, changing the alcohol during the first one to two days of storage is also recommended because samples release water and progressively dilute the preservative (Kilpatrick 2002; Nagy 2010).

Ethanol is flammable and considered hazardous.

- Researchers should avoid using distilled alcoholic beverages because they may have alcohol concentrations as low as 35%.

- Undiluted rectified spirits or neutral spirits (e.g., Everclear, Crystal Clear, Primasprit, Spirytus) is highly concentrated (95–96%) but should be avoided because it includes denaturing chemicals.

- Denatured alcohol (i.e., methylated spirits), widely used for industrial purposes, is made of 70–99% ethanol but contains additives that

make it non-consumable for humans (e.g., methyl ethyl ketone, also known as MEK) and thus should also be avoided (Post et al. 1993; Dillon et al. 1996).

DMSO

• DMSO is used as a preservative in many different aqueous solutions. One of the most commonly used formulations is a salt-saturated solution of DMSO (20% DMSO, 0.25 M ethylenediaminetetraacetic acid [EDTA], sodium chloride [NaCl] saturated, pH 7.5; Seutin et al. 1991, Nagy 2010), which is a cost-effective method for preserving and transporting genetic samples from remote locations, but the long-term effects on sample quality are still unknown (Williams 2007).

- DMSO solution has the advantage of being stable at room temperature, non-toxic and non-flammable. It can be carried on airplanes without a permit. However, it is photosensitive and can easily penetrate skin.

RNAlater

RNAlater solution and a similar product, AllprotectH Tissue Reagent (Qiagen, Valencia, CA), are used in sample collecting for the stabilization of DNA, RNA, and protein in tissues. Although these solutions minimize the need to immediately process and/or freeze tissue samples, material preserved in RNAlater and AllprotectH buffers cannot be stored at room temperature indefinitely if genetic samples are to remain viable.

Preserving Tissue Quality

• Minimize exposure to heat, light, moisture, and other chemicals
• Maintain cold chain if samples are frozen

Discipline Specific Information

Herpetology

• Liver and skeletal muscle are perhaps the most commonly sampled tissues

- A small incision can allow researchers to push the liver out, causing minimal damage to specimens being used for morphological

study.

- Bile salt can contaminate this organ and affect tissue stability, so tissue should be preserved as soon as possible and the gallbladder

avoided (Dessauer et al. 1990).

• Muscle can be dissected from the thigh on one side, leaving the remaining side intact for morphology, but it has been reported that yields are small due to tough fibers (Gamble 2014; Wong et al. 2012).
• Non-lethal sampling methods may include (Ezaz et al. 2009; Gamble 2014; Mollard 2018; Mollard et al. 2018):

- Biopsies

- Blood draws

- Feces collection

- Skin swabs (often used to test for chytrid fungus infection)

- Sperm or spawn collection (hormonally-induced)

- Toe clips

- Tail clips

Ichthyology

• Take tissue clips from right side (as for all destructive samples)
• Fin
• Muscle (hypaxial)
• Gill
• Whole animal (e.g. larval fish)
• Eye (especially from larvae), from right side

Mammalogy

Camacho-Sanchez (2013) found that rat (Rattus rattus) liver yielded the best RNA and DNA quality when compared to blood, brain, ear clips, muscle, and tail tips.

References

Nagy, Z.T. A hands-on overview of tissue preservation methods for molecular genetic analyses. Gamble, T. Collecting and Preserving Genetic Material for Herpetological Collection