Urine sampling protocol

The methods described here are currently used in Tuanan and other field projects, if you have any questions, please email Dr. Erin Vogel (erin.vogel*rutgers.edu) or Dr. Wendy Erb (wendy.erb*rutgers.edu).

Urine can be collected in three ways

  1. By using clean plastic and a catchpole to collect urine midstream as the animal urinates (note, plastic can be washed and reused)
  2. By placing a clean plastic sheet underneath a urinating animal or under an animal in a nest
  3. By pipetting directly from vegetation when uncontaminated by the urine of another animal (note if the leaves are wet from rain if this method is used).

NOTE: Collect as much urine as possible! What may seem like a miniscule amount of urine in the forest might actually be 400μL or 500μL, enough for two filter cards or tubes! Basically, you should leave the plastic or leaf dry…you can then later decide if have enough to discard

At the time of collection in the forest

- Record the name of the orangutan, date, and time on the side of the plastic tube using a permanent marker. Please write the date in the format 1 Nov 07 (not 1/11/07 or 11/1/07) and record time in 24 hour format (e.g., record 1:30 pm as 13:30). First morning voids are the best so please try to get these.
- If there is sufficient sample volume (> 0.5ml), use a urinalysis strip to test for the presence of ketones. Make sure the strip being used is not expired. Also, it is very important to keep the Chemstrip vials in silica gel so the batch does not get wet or moist from humidity. Thus, we recommend making urine collection “kits the night before going into the forest by placing 1-2 Chemstrips into the 15ml Falcon collection tube you will use. The strip can be dipped directly in the urine sample. Remove immediately, place on a clean surface, and read in 60 -90 seconds by comparing the strip colors against those on the side of the tube. Please take care to keep unused strips dry by closing container tightly. Record results in the datasheet while you are in the forest.
- Collect up to 15ml (the total volume of a 15ml Falcon tube). Then place tube in cold thermos.


At the end of the day / in camp

- Small amounts of feces and/or dirt will not affect our results. However, if you see that there is a lot of debris in the tube, please remove it with a plastic pipette. Also note if there is feces or dirt in the sample on the data sheet.
- An aliquoting order exists in order to prioritize urine allocation. With the sample, follow the procedure below:

  1. If you have a specific gravity meter, take specific gravity using 100 ul of sample, you can then pipet the sample back up to use on the filter paper (DO NOT DISCARD THE SAMPLE) – remember to clean off the SG meter with a tissue between samples.
  2. Prepare 2 x 200μL filter cards
  3. Prepare 5 x 200μL vials (put in 0.5ml tubes that we have)
  4. Prepare 2 x 200μL filter cards
  5. Prepare 4 x 1,000μL vials (put into the 2.0mL screw top tubes)
  6. Prepare additional 1,000μL vials and filter paper cards with any remaining urine

- If there is sufficient sample volume after following steps 1-4, use another urinalysis strip to test for the presence of ketones. Make sure the strip being used is not expired! Record results in the datasheet.



If using Whatman Protein Saver Cards, cut the filter out of the envelope but be sure not to touch it with your bare hands – WEAR GLOVES. Cut the card into approximately 2cm x 2cm squares to aliquot urine on (about 8 squares). If using rolled filter paper, cut squares with similar dimensions. NOTE: Always record how much urine is applied to each card in both the data sheet and on the card (should be 200μL unless a special circumstance)

  1. Prepare a “boat” from aluminum foil, by cutting a square and folding up all 4 sides to a height of ~1cm. This prevents the filter papers from sliding off during transfer and storage. Ensure the square is large enough to hold the number of filter papers being prepared. Prior to pipetting urine, record the following information on each filter square as seen below PENCIL: Sample # and a unique letter for each card (starting with “A”, followed by “B”, etc.)
    OU Name
    Volume (μL) (should be 200μL unless under special circumstances)
  2. Place the filter papers in the foil, taking care that the papers are not touching each other.
  3. Use the micropipetter to apply 200μL of urine to the center of the filter paper square. We have found that you should not exceed 200μL per card as the cards get too saturated.
  4. When finished applying urine to all filter cards, place the boat in the drying container, seal the container, and allow the urine to dry onto the filter paper for 24-48 hours.
  5. When completely dry, place each filter square into its own sleeve in a slide sheet. Place the filled slide sheets into the designated container (this is separate from the one used for drying filter cards).
  6. Store filled slide sheets in a designated airtight storage container with silica gel, preferably in an AC room.



  1. Use the micropipetter to aliquot the desired amount of urine. Be sure you set the dial on the micropipetter and select the appropriate pipette tip size. Use the 0.5mL tubes for the 200μL samples, and 2 mL tubes for the 1,000μL samples and follow the protocol above for aliquot amounts (see “At the end of the day / in camp”). Any extra urine can be stored as 200μL aliquots on filter paper and 1000μL aliquots in tubes. Leave a small space at the top of the tube to prevent the tube cap from bursting once the sample is frozen (hence 200μL in 0.5ml tubes and 1000μL in 2.0mL tubes).
  2. Using the label maker (best) or a permanent marker, record the sample # and letter on the cap and on the side of the test tube. The sample numbers should represent a single void and each void should be assigned a new number (should be in consecutive order). DO NOT COMBINE samples from different voids, even if they’re from the same OU on the same day!! For each sample #, a unique letter should be assigned to each vial, beginning with A (e.g., #421A, #421B, etc.). Note that filter card letter labels and vial letters can be redundant (e.g., there may be a filter sample #421A and a vial sample #421A). Record the volume of the sample on the side of the tube as well. Do not date these tubes (this is very important – just number and volume). Please make sure the number on the tube and the number in the data log book match.
  3. Place the samples in the next available empty cells in the sample storage box located in the freezer and immediately store the samples in the freezer. When starting a new box, write Tuanan followed by the next consecutive number that follows the previous box (e.g., Tuanan 12 follows Tuanan 11).
  4. Promptly store the sample in the freezer and check that the freezer is working properly. If not, make plans to evacuate all samples to the backup freezer. If the freezer is found broken/not working sufficiently, and the samples have thawed, record this incident of affected samples.
  5. Rinse the 15 mL tube and place in tub labeled bottles that need to be cleaned. These bottles should later be washed well, the ink removed using alcohol wipes, and can be used again.



- Record the basic information in the sample log book in the lab (sample #, OU name, time of collection, person who collected, number of tubes, and number of filter cards/papers prepared).
- Remember to completely fill out the data sheets (copies are provided in the office alongside the other data sheets). Fill out EVERYTHING on the data sheets including:

  1. Sample # – unique identifying number, assigned in consecutive order
  2. Date – date sample collected
  3. Time – time sample collected (use military time)
  4. Collected by – name of individual who collected the sample
  5. OU Name – write out the name of the individual sampled
  6. Age-Sex – AF (adult female), SAF (Subadult female), JF/JM (Immature female or male), UM (unflanged male), FM (flanged male)
  7. Mating observed? – Was the individual observed to mate on this day
  8. Association – how many and what types of individuals were in association with the animal at the time of collection
  9. How sample was collected – from plastic, leaves, or both
  10. FP# - the number of filter paper circles prepared (if any). Please see attached instructions for preparation of filter paper samples
  11. Conditions at time of collection – raining, wet, dry, etc.
  12. Notes – any other relevant information. Long notes can be continued on the next line.
  13. Results of Chemstrip – record results from all tests, indicating time of testing
  14. Details of samples prepared – Sample letters and quantities for both filter cards and vials, including time that samples were stored in freezer
  15. Box# – the box ID in which the urine tubes are stored
  16. Collected by – individual who collected the sample (this includes those also following, if A collects the urine while B is also there, the data sheet should state A first, B second)


Silica gel

The use of silica gel desiccant is essential to keep filter paper samples dry and free from mold. The crystals absorb moisture but eventually reach a capacity in which they cannot absorb any more moisture. Therefore it is important to check the silica periodically and replace as needed. We have provided you with “color-indicating” silica. These crystals are blue when active. Once they have absorbed all they can absorb, they turn pink and should be reactivated and/or replaced. To reactivate the gel, use the designated pan in the office to gently reheat the gel, stirring regularly, until all crystals have turned blue again. The gel can be reused indefinitely. Please take care when rewarming or disposing of silica gel. It is poisonous if ingested by humans or other animals.