Hi IImari,

Studies have consistenly shown that the use of cotton-based materials, such as the salivette, affects the realibility of nearly all salivary analytes, and often quite substantial. I cite several of those studies in my review on the usefulness of salivary amylase (Bosch et al. 2011, Psychoneuroendocrinology), but there are much in the literature. All showing the same thing. So, despite their tremendous popularity the data are strongly biased against using such devices.

There is now a cortisol salivette on the market that uses a synthetic material that seems to give more reliable results for salivary cortisol, but to my knowledge it has not been well-tested for other salivary components. Usage would appear at your own risk therefore.

The most reliable, and the cheapest, way of collecting saliva is to have participants spit in a (polypropylene!) tube of closeable cup. Look up a catalogue of companies that sell those, and you will find many shapes and sizes that are practical. Amongst saliva researchers that approach is considered the gold standard also because it allows reliable measurement of saliva flow rate. Some researchers fear that participants may find this spitting procedure embarressing, but this idea has not been supported by the data (see my review for relevant references).

I hope this helps!

Jos

Hi Ilmari,

You are asking about Testosterone.  The issues with T can be different than Cortisol.  We tried salivettes and the results were not as reliable as with simple passive drool into plastic cups, disposable pipette into test tube. I think that our T RIAs were less affected by salivettes than ELISAs.  But collection methods depend on context.  In Dominica we never had any issues with children & parents (& grandparents etc.) passive drooling.  For infants we used dental cotton pressed out in a syringe, but never looked at T from them.  I never wanted the risk of a kid choking on a salivette.  On the other hand, we recently started a study of law school students, who could not handle the trauma of passive drooling.  Go figure.   

I don't envy researchers who can't collect saliva directly from participants.   It is critical to know precise time of collection (and wake-up), and flow rate.

Hi Ilmari,

definitely the commercial sampling devices affect sex hormones levels determined in saliva, spitting is the "gold standard" - see papers from prof. Celec's group: 

http://www-ncbi-nlm-nih-gov.ezp-prod1.hul.harvard.edu/pubmed/22579929

http://www-ncbi-nlm-nih-gov.ezp-prod1.hul.harvard.edu/pubmed/24051109

katka

Hi

I was wonder whether if an extraction of the saliva could reduce the high variability in the EIA? Bests, Virginie

Hi Ilmari,

The cotton-based salivette (Sarstedt) will certainly interfer with testosterone measures. As stated above, synthetic swabs (Salimetrics https://www.salimetrics.com/collection-system/adult-oral-swab) should not cause interference in testosterone EIA testing. Passive drool (whole saliva) is the "gold standard", but I can assure you from my years working at Salimetrics that their swab product is validated versus passive drool as part of the QC process.  You could also ask if they would share their validation data.

Products are available in Europe through their UK branch. Ask for samples!

Good luck!

Mary

Hello! Thanks for all! You seem to be all agreeing, so I might have to change the methods. It appears to me that the passive drool collecting methods are more expensive than the Salivettes I have bought before (100 dollars for 50). Do you have any comments on the price?

Also, an additional question, which might be a basic one: how do you collect and keep your samples after collection? For instance: if the subject collects data at their home, is it OK if they try to keep the samples cool (fridge temperature) for some days and then provide them to me in normal bags (thus, they might get warmer than 4 C at some point).

I found this description - do you agree with it? So, up to 4 months a normal freezer is OK and beyond that, a special freezer is needed?

"Sample Handling and Processing (As described in the Saliva Collection Handbook):

• Immediately after collection, freeze samples at or below -20°C. If freezing is not possible, refrigerate immediately at 4°C and maintain at this temperature for no longer than necessary (ideally less than 2 hours) before freezing at or below -20°C (temperature of a regular household freezer).

• Samples stored for more than 4 months should be frozen at -80°C.

• Freeze-thaw cycles should be minimized for some analytes. It is critical that storage conditions are researched prior to initiation of sample collection."

For -80 C I would need to buy a special freezer. The cheapest I could find is 3800 euros for 320 liters (or dm3) - does this sound like a reasonable price? Does anyone have experience about special freezers?

Hi Ilmari,

If you are going to store saliva unfrozen then you will have problems.  I used sodium azide as a preservative for inhibiting bacterial growth and enzyme activity, but always got samples cold ASAP.  I would be very careful using sodium azide with tubes provided to subjects because there would be a risk of poisoning.  If they follow directions and drool into cup then pipet into tube there is no risk.  But....

Here is a summary of the methods I used.  

STANDARD RECIPE

You will need:

polystyrene tubes with caps, sterile,

Fisher: O.D. x L: 13 x 100mm; Closure: With screw cap; Sterile; Bag of 125

14-959-38C

Bd Vacutainer Labware Medical

No.:352027 Case of 1000 for $351.52

Sodium azide (get it from someone locally if you can; you will only need a pinch) Make up a one litre bottle (strong screw cap) of solution (or two .5 litre bottles, so as to put all your eggs in one basket -- also bring one bottle of distilled water for rinsing), mark it POISON with a sharpie, skull and cross bones etc. Solution that I use is 5 grams per liter.

No need to get any more

Sterile double-distilled water (for use in making azide solution, and rinsing syringe pipet after dispensing azide solution)

Free, just need a few bottles for field

In the field I shoot .1 ml of the azide solution into each tube with the syringe dispenser (decap the tubes, put them into a foam rack, pipet azide solution into each tube, put caps back on tubes -- I usually do a large batch at one go).

I use the foam racks to carry the tubes around during collection. I have a medium sized, long bag with a zipper top into which I put a piece of cardboard to have a firm bottom. If you collect 2.5 ml of saliva (I usually try for about 3-4 ml so as to have plenty for different assays), that will give you about a .2 g/l mix of azide to saliva, which has proven effective in preventing most degradation. Keep saliva out of the sun if possible. Time the collection and record how long it took and how much saliva approx. was deposited in the plastic cup.

Collection bag (carries foam rack with tubes, gum, pipets, plastic cups, clipboard with data collection sheets, small camera, sharpie pens, etc.)

Wrigley's spearmint gum (a quarter stick will usually do fine; let subject chew it and swallow sugar for about 1 minute, then begin expectorating into plastic cup, preferably with head canted slightly down, so that little or no saliva is swallowed and a flow rate can then be reasonably estimated) -- but the important thing is not to freak anyone out, do what is comfortable for them, and try to be as reinforcing as possible. I sometimes chew gum while I dispense it to subjects, so that they see me doing the same thing, i.e., the gum is not poison. If you are dealing exclusively with adults in resting states, you may wish to forego using gum or any stimulant.

Gum is about $3 for ten pack 5 stick packages = 50 sticks, I break in four, so $15 per thousand saliva samples.

Syringe dispenser (I have a Nichiryo model 8100) to pipet azide solution into tubes -- they are a bit pricey ($200), and you could use a disposable plastic pipet in a pinch -- the amount of azide solution does not have to be super precise, although remember that you are in effect diluting the saliva by a set amount (approx 3%), which should be constant.

plastic cups

About $10 box of 1,000

disposable plastic pipets (to move saliva from cups into tubes)

Fisher [disposable pipets]

Graduated; Capacity: 7.5mL; Length: 6 in.

13-711-9D Case of 10 PK (5000) for $344.92

Tube labels -- stick on (if you can computer print name, date etc. in field that is great; otherwise handprint clearly-large, and number your tubes and data collection sheets (see below) so that the numbers match the tube with the collection sheet data.

About $10 box of 1000

A data collection sheet to record info about sleep (wake up time), time of collection, food, activities, health condition, caffeine, tobacco or alcohol use, etc.

Foam test tube racks to hold tubes while you pipet azide solution into them, and for carrying tubes to and from collection sites.

Fisher: [one case foam racks for each site]

Holds 50; For 10mm, 12mm or 13mm diameter Tubes

05-664-15A Case of 5 PK for $270.00

Parafilm (wax film) to wrap filled test tubes (it comes it big rolls; you cut about 1 inch pieces, take off the paper, then hold wax on the tube with one thumb, then stretch with your other hand and wrap around the tube. The idea is to get a secure seal around the tube cap. Do this after you have returned from collection, and gotten your numbers all straight). Use scissors to cut pieces; I usually cut a ton and stash them in tupperware containers, but experiment first to get comfortable with the size, stretch, etc. Practice at least 20 or 30 empty tubes before you wrap a valuable filled tube. If you do not get a good seal, the caps may blow off during airtravel.

Don’t need with screw caps unless you are anal about hygiene.

Tupperware containers to use for storage of caps, parafilm, etc. during azide pipetting

Thick freezer zip lock bags medium large to stash filled test tubes (150 tubes per bag)

About $10 per 100 bags.

rubber bands to bundle batches of ten test tubes for placement in zip lock bags

Sharpie pens of various colors

Tubes, pipets, azide, parafilm, bottles, syringe dispenser, foam tube racks, etc. can all be obtained from Fisher Scientific. I get the cups in the field as they are bulky to bother travelling with.

Make sure you have a letter from the Minister of Health/medicine to bring all this stuff in; test tubes etc. make Customs officials nervous, esp in drug lands...

Procedure is a bit different for infants, but I gather you are not interested in them.

The lab work is much more complex and sensitive. We used an ATX 180 that can process 180 samples per hour, do multiple assays in one run (e.g., estrogen, estradiol, cortisol, and testosterone), straight from the original sample tubes. The lab was at the University of Michigan Hospitals -- I worked with Barry England -- I now work with Michael Muehlenbein (Indiana) and Pablo Nepomnaschy (Simon Fraser).

The best way to learn all this is to see it in action. Next time I am in Dominica I will let you know and you are welcome to drop by and check it out. If you get a big grant and want to fly me in for a couple weeks I might be able to do that if the timing was right...

More on this if I think of something I forgot.

END OF STANDARD RECIPE

thanks a lot! I already edited my question a little as I found some more information.

Do you think sodium azide is needed if the samples are frozen (-20 or -80 C)?

Check the data sheets for the assay kits you will be using prior to collecting saliva in sodium azide tubes! (I know for a fact that sodium azide will interfer with Salimetrics salivary assays.) Freezing samples soon after collection to minimize bacterial growth is usually the preferred storage method.

You can collect passive drool in the salivette tubes: remove the plastic insert and cotton swab. The large tube and cap will work fine for passive drool (whole saliva) collection. I frown on the use of gum since it has the potential of changing the salivary pH thereby causing assay interference. Making sure subjects are well-hydrated and/or drink water 10 minutes prior to start of collection should provide you with an adequate sample. Collecting 3-4 mLs can be difficult for some people and take >15 minutes. 

Samples can be aliquoted into smaller vials in your lab. Ideally this should happen before they are frozen for the first time, but if that's not possible, aliquot the sample after the first freeze/thaw cycle. Use cryolabels so that labels will not fall off the tubes. It's always a good idea to write the information on the tube using a permanent marker before placing the label on the tube. Good insurance should the labels fall off upon thawing.

A non-cycling (manual defrost) freezer should be used for long-term sample storage. -80C freezers are very expensive, but you can get by with a commercial grade -20C freezer for storage up to 1 year. All dependent on how much money you can spare.

Good luck!

Mark, thanks a lot for the very detailed answer! actually, there are some exchange grants included in some grants so I'll have to see about the Dominica-project in the future! :)

Mary's points are great.  Be careful if you use gum or any other stimulant to minimize potential contaminants.   Mark tubes directly in addition to labels. 

Note that anthropological field conditions are often different from folks living in apartments in Helsinki etc.  

One thing I have to laugh about is how 'wimpy' folks in the Western world are about providing saliva.  Karen Strier once commented after I gave a talk that when she and other grad students at harvard had to provide saliva samples it was almost torture; whereas the kids in my study had no issues providing several samples every day, day after day after day for years.  Go figure.  

Dear Ilmary,

I have four comments that you may find useful:

Passive drool; that term is a bit of an umbrella  and is used for both actual passive drool and the spitting method. Both are equally reliable, but the latter, I find, tends to be easier and more pleasant for participants. For a detailed protocol (with pictures) see Navazesh 2008 J Am Dent Assoc.

Second, I was surprised by your comment that "passive drool collecting methods are more expensive". That is definitely not the case: many companies sell closable small buckets or centrifugable tubes that are both good and very cheap (I am talking cents per piece). Always use polypropylene, never polysterene.

Third, some saliva components are very stable at room temperature or 4C, but some are not. There is data on that, but it is worth considering (and very straightforward)  to test that in your own lab: Compare samples kept under condition X or Y before freezing with immediately frozen samples. It may also deliver you a nice and useful publication!

The real trouble may however occur after thawing the samples, because freezing/thawing breaks up buccal cells and bacteria that contain most of the degrading enzymes. Most oral biology labs therefore centrifuge samples before freezing (to remove cells) and only freeze the supernatant. Additionally consider keeping the samples cold/on ice after thawing. Azide appears only useful with some saliva components: check out the work of prof Helmerhost at Boston University, who is the expert on this topic

Hope this is useful!

Jos.

Note the brain fart mistake in my list:  as Jos notes, use polypropylene tubes, not polystyrene.

Thank you all for very useful answers! Actually, even the most simple advice can be useful when you (meaning I) haven't done something before.

Jos,

one more question:

so you suggest centrifuging the samples before freezing, and minimize freezing cycles (also the manuals said so). So, are you saying the samples should be centrifuged as soon as possible? How about samples that are impossible to be centrifuged right away - should they be freezed and de-freezed later for centrifugation and freezed again for a longer storing? So, I don't know which one is more serious for the samples. I guess, it would be best to freeze them as soon as possible, and all else comes after that? The samples are not analyzed in the same place where they are freezed in the first place.

In our Lab we tested the differences between passive drool and cotton rolls like these ones (http://www.servi-dent.com/wp-content/uploads/2013/09/Cotton-Rolls.jpg).

We found no statistically significant differences regarding total volume, total protein content and SDS-PAGE profile.