Alberts Laboratory Procedures for Fecal Genotyping and Paternity Analyses

Alberts Lab Fecal Protocol, 30 Oct 2012


Prepared by Jacob Gordon, with significant inspiration from the Altmann Lab Protocol


Samples are collected ad libidum from free-living baboons living in the Amboseli basin of Kenya. When a known animal is observed defecating, we collect the freshly-dropped sample within minutes of its deposit. Unlike the samples collected for hormone analysis, the fecal sample is not homogenized prior to collection. Instead, a small portion is removed from the leading edge of the fecal bolus (if possible) and placed in a plastic 20-mL vial (Evergreen Scientific 220-3519- 080; caps 300-3532-G20) prefilled with 95% ethanol (EtOH). The sample if broken up to saturate it with EtOH and the vial is identified with the baboon’s name (ID), the sample date, and time of collection. Samples are stored in an evaporatively cooled hut (a “charcoal fridge”), which keeps temperatures 15°C-25°C (~daily min, max). A list of all samples, including the baboon id, the date and time of collection, and any additional notes about the sample, accompanies the samples when they are eventually sent to Nairobi.

Unlike the hormone fecal samples, the genetics samples do not undergo any processing in Nairobi. They are stored there and eventually shipped to Duke University.


Upon arrival at Duke, the shipped feces are sorted and inventoried by at least one technician (ideally two, to screen for typos). Tubes are stored in freezers at -85°C. A record of each sample, including its name, collection date, time of collection, and location in the freezers, is maintained in our online database, Babase. The database also assigns a unique Tissue ID number (TID), which is referred to below.


Until ~late 2009, fecal DNA extractions were performed using the Qiagen DNA Stool Mini Kit (cat. #51504). Since then, extractions have been done with a protocol from Qiagen using the Qiagen QIAxtractor, modified somewhat to accommodate the peculiarities of fecal DNA extraction. That protocol is outlined below:

·  Preparation

o  Turn on 70°C water bath (not necessary if you plan to stop at end of fecal transfer)

o  For cleaning spatulas (more info below), set up 4 50-mL falcon tubes:

§  Two with 40-45 mL dH2O

§  One with 40-45 mL bleach

§  One with 40-45 mL 95% EtOH

§  Line up tubes next to each other in the following order: water, bleach, water, EtOH

o  In a biosafety cabinet, ignite a Bunsen burner, EtOH burner, or other open flame

o  Check the bottle of ASL buffer for precipitation. If precipitate is present, place in the heating-up water bath and allow the heat to dissolve the precipitate. (not necessary if you plan to stop at end of fecal transfer)

o  Don a gown, hairnet, and facemask

·  Fecal Transfer

o  Using metal spatulas, transfer fecal samples into 2-mL round-bottom centrifuge tubes up to the 0.5 mL mark.

§  If it's possible to identify the frontmost part of the fecal bolus, try to collect from there. We expect a higher amount of baboon epithelial cells there.

§  Be careful to not go above 0.5 ml; if anything, err on the side of caution and go less.

§  We prefer 16 samples per QIAxtractor run

§  Use a clean spatula for each sample

§  To clean spatulas (be sure to do this with spatulas at beginning of procedure, before doing any transfers) :

·  After using a spatula, place it in first water tube

·  When 4-6 spatulas accumulate in first tube, move them to the bleach tube and allow them to soak for at least a few seconds. It's okay to continue transfers for 4-6 more samples and move spatulas out of bleach when the next group is ready to go in.

·  Remove from bleach soak and move spatulas to the second water tube to rinse bleach. Extended soak is not necessary, just be sure rinse does occur.

·  Move spatulas from second water tube into EtOH. Extended soak not necessary.

·  After EtOH tube, wave EtOH-soaked spatulas over open flame and allow ignited EtOH to clean and dry spatulas. Don't get burned!

·  Lay spatulas across a lab marker or some way such that the ends that will be touching feces aren't touching anything.

·  Water tubes should be emptied and refilled after each extraction, but bleach and EtOH tubes can usually be reused for 2-3 total extractions. When in doubt, throw it out.

o  If necessary, it's okay to pause the procedure at this point

·  Lysis

o  If you paused the procedure at fecal transfer and are resuming now, turn on 70°C water bath, and if precipitation in ASL buffer has occurred, heat it up as described above

o  To each tube, add ASL buffer (make sure precipitate is not present in buffer first) up to near the top, ~1600 µl.

§  This feces:buffer ratio is fairly sensitive. As mentioned above, it's better to err on the side of having less feces; DNA yield decreases sharply as the amount of feces increases, presumably due to inhibitors in the feces.

§  It's important to not stop at this point. If precipitate from ASL forms again, DNA will likely precipitate with it and may not return to solution if heat is re-applied. We have heard from collaborators that overnight incubation at 37°C (which should be sufficient to prevent precipitation) increases fecal DNA yield in other kits, but we haven't tried it enough with the QIAxtractor to know.

o  Make sure all caps are securely and firmly closed, and vortex all tubes for at least 15 minutes, or until all samples are homogenized.

o  Incubate homogenized samples in 70°C water bath for at least 5 minutes

·  Centrifugation/Supernatant Collection

o  Centrifuge tubes at maximum speed (~14000 RPM, ~18000 × g) for 10 minutes

§  Spinning less time than this tends to not sufficiently pull down invisible contaminants that often cause problems later on in the QIAxtractor

§  Spinning more time than this may (we haven't done any experimentation on this question) pull some DNA into the pellet

o  Transfer supernatant to separate wells of a QIAxtractor lysis plate, 200 ul supernatant/well. When doing 16 samples, we use the following layout:

Samp 1 / Samp 1 / Samp 1 / Samp 1 / Samp 1 / Samp 1 / Samp 9 / Samp 9 / Samp 9 / Samp 9 / Samp 9 / Samp 9
Samp 2 / Samp 2 / Samp 2 / Samp 2 / Samp 2 / Samp 2 / Samp 10 / Samp 10 / Samp 10 / Samp 10 / Samp 10 / Samp 10
Samp 3 / Samp 3 / Samp 3 / Samp 3 / Samp 3 / Samp 3 / Samp 11 / Samp 11 / Samp 11 / Samp 11 / Samp 11 / Samp 11
Samp 4 / Samp 4 / Samp 4 / Samp 4 / Samp 4 / Samp 4 / Samp 12 / Samp 12 / Samp 12 / Samp 12 / Samp 12 / Samp 12
Samp 5 / Samp 5 / Samp 5 / Samp 5 / Samp 5 / Samp 5 / Samp 13 / Samp 13 / Samp 13 / Samp 13 / Samp 13 / Samp 13
Samp 6 / Samp 6 / Samp 6 / Samp 6 / Samp 6 / Samp 6 / Samp 14 / Samp 14 / Samp 14 / Samp 14 / Samp 14 / Samp 14
Samp 7 / Samp 7 / Samp 7 / Samp 7 / Samp 7 / Samp 7 / Samp 15 / Samp 15 / Samp 15 / Samp 15 / Samp 15 / Samp 15
Samp 8 / Samp 8 / Samp 8 / Samp 8 / Samp 8 / Samp 8 / Samp 16 / Samp 16 / Samp 16 / Samp 16 / Samp 16 / Samp 16

§  Take extreme care to avoid any insoluble matter when transferring

·  Fecal lysates often have a mucusy material that usually rests at the top of the pellet; this matter often causes membrane clogs in the QIAxtractor

·  There are other particulates, often invisible at this stage, that can cause clogs in QIAxtractor, so keep pipet tip as far from pellet as possible

·  QIAxtractor

o  Bring plate to the QIAxtractor

o  For DNA capture, we find that the VX capture plate (Qiagen cat. #950921) gives much, much better yields of endogenous DNA than the DX plate (#950911).

§  Aside from the VX capture plate, use only DX reagents (they're the same price or cheaper)

·  We've tried using all VX reagents, but saw no notable improvement in DNA yield.

o  Reagent Prep

§  DXW, DXF, and DXE (we use autoclaved ddH2O instead of DXE) can be poured into respective reagent troughs far in advance, like during sample vortexing, heating, or centrifugation. For each trough, add the volume directed by the software wizard.

§  DXB binding additive (refrigerated white solid) needs to be added to buffer DXB, but once added they only stay stable for a month at most. If we expect to do a lot of extractions, we mix one vial of binding additive with one bottle of DXB and use the quantity that the software's wizard directs. If we expect few extractions in the near future, the QIAxtractor manual has an appendix showing precise amounts (in mg) of additive to add to desired amount of DXB, and we mix only the amount we need

·  Binding additive needs to be refrigerated when solid, but once added to buffer, do not refrigerate buffer DXB. Cold storage causes precipitation of the binding agent, so store at room temperature.

·  Mixed DXB is photosensitive, so don't add DXB until just before DXL and before starting cycle

§  DXL: dilute digest enzyme (DX) 1:10 in DXL buffer to a final volume of whatever the software's wizard tells you it needs.

·  Once diluted, the enzyme doesn't stay stable for long, so do this just before starting the cycle

o  Dealing with clogs

§  There are two steps in the QIAxtractor protocol where the program waits for confirmation from the user before it will proceed onward. Before allowing the procedure to continue, it's important to look at the wells and make sure no wells are clogged—they should be dry. If fluid is still present in any wells, open the lid and try pipetting the residual liquid up and down a few times to dislodge whatever clog is present. If the clog is dislodged, you'll see the movement of fluid pretty soon. If no movement occurs, manually aspirate fluid as best you can without puncturing the membrane, and then allow the procedure to move onward.

§  If any clogs are visible at the second checkpoint, check on the plate periodically for the rest of the cycle to make sure reagent added to the clogged well(s) doesn't overflow into adjacent wells.

·  After QIAxtractor

o  Elution tubes from QIAxtractor are not good for long-term storage. No matter how tightly you place the caps on, cold storage causes the caps to pop and the fluid sublimates over time in the freezer. Storage for a night or two isn't likely to be harmful, but it's best to transfer eluted DNA's to screwcap tubes as soon as possible.

Each extracted DNA sample is assigned a DNA ID number (DID), and we record basic information about the sample, including name, extraction date, technician(s) involved with extraction, and the TID of the tissue source. All this information is stored in Babase.

After extraction and transfer to screwcap tubes, DNA's are stored at -20°C.


Fecal DNA extracts are expected to have large amounts of non-baboon DNA (especially bacterial), so spectrophotometric quantification (e.g. Nanodrop) is not very helpful. Instead, we quantify our fecal DNA extracts using a qPCR assay for mammalian c-myc, described in Morin et al, Molecular Ecology 2001. We order probe and primers from IDT, and use the Qiagen Quantifast Probe PCR Kit (cat. #204254), with the following recipe, from the kit's manual:

1X 50X 100X

Water 8 ml 400 ml 800 ml

2X Mastermix 12.5 ml 625 ml 1250 ml

10mM Forward Primer 1.0 ml 50 ml 100 ml

10mM Reverse Primer 1.0 ml 50 ml 100 ml

10mM Probe 0.5ml 25 ml 50 ml

Template 2.0ml

Total Volume 25ml

PCR conditions:

1 ´ 95°C, 10 mins

40 ´ 95°C for 15 sec and 59°C for 30 sec

For our standard, we use ABI's TaqMan Control Genomic DNA (cat. #4312660), serially diluted 1:4, from 10 ng/mL (stock) down to 9.8 pg/mL.

Concentration according to qPCR is recorded in Babase, and should be imported to Babase as soon as possible after qPCR is complete.


To determine an individual's genotype, we PCR at 14 distinct microsatellite loci, using Qiagen's Multiplex PCR Kit (cat. # 206143, 206145). Qiagen's protocol is for 50-mL reactions; we scale down to 10 mL, as shown below:


Water To 10 mL

2X Mastermix 5 mL

2 mM Primer Mix* 1 mL

DNA 1-2 mL

*Primer Mix is a mixture of forward and reverse primers for all loci being assayed in the reaction. Each individual primer has a 2 mM concentration, so that the final concentration in the PCR of each primer will be 0.2 mM.

Primer Info:

Locus / Size Range / DNA/well (mL) / Fwd Primer / Rev Primer

Using IDT and ABI to generate our oligos, we use the above primer pairs, with one of the primers labeled with the dyes FAM, HEX, or NED. Each PCR well usually uses primer multiplexes of 3-4 pairs. In designing multiplexes, we take care to not use the same dye for oligos of overlapping size range.