This simple protocol provides the necessary basics for hookworm incubation. Anyone wanting more detail should read Alana's very detailed NA incubation method.
Preparing the mixture
If you are using a donor, have them do this step if they don’t live nearby and if it is going to take over 20 minutes to get the sample to you.
- wide mouth container (urine container, mason jar, etc.) with screw tight lid
- barrier gloves (latex, nitrile, etc.)
- gallon ziplock bag
- waxed disposable plates (for a work surface and resting place for contaminated instruments)
- plastic knife
- vermiculite (with no added material)
- activated charcoal (capsules are fine)
- distilled water (filtered or dechlorinated tap water and bottled water are also suitable)
Take some feces from a hookworm host who has been infected for at least 6 weeks. (Eggs aren’t produced before this, and the quantity of eggs increases in the weeks following, up to a maximum at 20 weeks.) The feces used must also be fresh because hookworm eggs degenerate rapidly during the first month. 
Place the feces in a covered container. A large amount of feces is not necessary, a few heaped teaspoons will be plenty.
Add vermiculite (one part vermiculite to one part stool) and activated charcoal. I use one charcoal capsule, or two if I’m doing a big batch. The charcoal kills the smell and balances the pH (healthy stool is acidic, but a neutral pH of 6.0 or 7.0 is optimal for hatching hookworm eggs) and the vermiculite retains moisture so prevents the mixture drying out.
If necessary, add distilled water to get to a pudding consistency.
Use a knife or fork to mash any large chunks and stir until smooth.
Wash outside of container with soap and water or alcohol swabs.
Clean up by placing supplies in ziplock bags and into freezer overnight to kill any residual material.
Now you are ready to incubate.
Supplies for the rest of the process
- wide mouth container (urine container, mason jar, etc.) with screw tight lid. It's also possible to use test tubes, or centrifuge tubes with conical bases placed in upturned egg cartons. In this case, the larvae collect conveniently in the tube’s conical base. 
- barrier gloves (latex, nitrile, etc.)
- gallon ziplock bag
- waxed disposable paper plates (for a work surface)
- glass microscope slides
- coffee filter paper (or flat, uncoloured paper towel)
- plastic spoon
- distilled water
- conical-bottomed container such as a wine glass, silicone snow cone cup, or glass test/centrifuge tube
- microscope (This needs to be capable of 100x magnification, preferably binocular compound and with a moveable stage. See: Microscope selection notes.)
- tape/ bandage
Cut a strip of filter paper to the size of the glass slide, leaving an area at one end free, so you can pick up the slide without touching the filter paper. If using centrifuge tubes, cut the paper slightly narrower than the slide to prevent it from touching the sides of the tube. (A wooden craft stick makes a suitable alternative to a glass slide if it is cut to size with shears so it fits inside the container. Old credit cards and sections of CD cases or takeaway food containers can also be used, and these will hold more stool than sticks or slides.)
Wet the paper and put it on the slide.
Put about half a teaspoon of the feces/vermiculite mixture on the middle of the slide. (This should yield at least 100 larvae.)
Add some water to the bottom of the jar. The mature larvae will wriggle into this water. (Note: use only enough water to wet the bottom of the filter paper, because you’ll have to search through all this water later when you look for larvae!)
If you have more prepared mixture, you can set up multiple jars, which is a good idea since some incubations can fail inexplicably. (See Possible reasons for failure.)
Let the container sit somewhere away from light for 1-2 weeks at room temperature. No incubator is needed if the temperature is 70ºF (21ºC) or above, and you can put it outside if the weather is warm. (Place a jar of coconut oil nearby. The oil should be liquid if warm enough.) If the ambient temperature is too low, place the container in an airing cupboard,  use an egg or reptile incubator,  a plant propagator, or a suitable container plus a heat source.  For more detail about incubation temperature, see Incubation temperature, below.
Keep the container covered to make sure no bugs can get inside. Mold may form due to the high moisture, but this doesn’t matter because it won’t affect the worms. The only problem I have with the mold is sometimes larvae get stuck to it when I’m trying to count them in the solution.
If you use a coffee filter or cloth over the top of the container instead of a lid you will get less mold but there's more chance of it drying out. If you use cloth you will need to mist daily. See Keeping the sample moist, below.
I have had the most failures due to forgetting about it and letting it dry out. NA larvae die in a matter of minutes once they become dry. 
Hookworm eggs do need air for development  but there is plenty of air for them in a jar, even when this is sealed with an airtight lid.  If using smaller containers, such as test tubes or urine sample bottles with airtight lids, it may help if the lids are removed briefly every day to provide an air change.
(NB. The presenter of this video says he sets the thermometer on his incubator to 99°F (37.°C), but this temperature is too high for hookworm incubation, and he does mention that the thermometer he is using may not be accurate. See Incubation temperature, below, for guidance regarding the best temperature to use.)
One of the benefits of using this method is that only L3 larvae can escape from the feces and be collected in the water.
After a week or two, pipette all the liquid from the bottom of the container and put this liquid into a conical-bottomed container such as a wine glass, silicone snow cone cup, or glass centrifuge tube. Let this sit for at least a few minutes so the larvae - which cannot swim - have time to sink and collect at the bottom.   The majority of the larvae will have settled in 2 or 3 hours,  and all of them by 24 hours.
An alternative to using a conical-bottomed container is to draw up the liquid into a transfer pipette and then stand this up in a container.
Pipette the water from the bottom of the wine glass, or alternative container, onto the middle of a glass slide. Squeezing the pipette bulb before it enters the liquid will prevent bubbles from stirring up the collected larvae. Some growers recommend pipetting from 2 to 4 mm above the bottom to minimise the number of dead larvae that are collected, and to save time at the next stage,  but others don't bother doing this, and just pipette right from the bottom. One grower swirls the larvae and then lets them settle for a few minutes, and recommends practicing this with fine dregs from tea leaves, or something similar, to watch how the swirling causes them to gather. 
Place the slide (without a cover slip) on the microscope's stage and set the magnification to 40x. Then, moving systematically up and down across the fluid on the slide, examine for larvae.
Some people may find it easier if a few small drops of fluid are placed on the slide rather than a single large drop.
It's easier to get small drops if you use a fixed-volume micropipette/minipipette. e.g., a 100 microlitre mini pipet such as this model which takes removable tips.
A single drop from a 100 microlitre micropipette may contain 100 larvae, but 10 larvae per drop is easier to count. If there are too many larvae, add more water to each drop, or divide each drop on the slide using a pipette tip, and then add water to these daughter droplets.
If you don’t get any larvae from the container, or don’t get enough larvae, add more room temperature distilled water to the bottom of the container. Repeat until the desired number of larvae are obtained.
I only use squirming larvae. I poke them with a mechanical pencil to see if they are alive (requires a steady hand and practice), but larvae that are not moving are not necessarily dead. They conserve energy until they find some skin, so motionless larvae that are intact may be viable. (I use 100x magnification for a closer look at individual larvae.) Scraping some skin cells from an arm and adding these to the water will often get the larvae moving, as will gently pushing them to the edge of the drop of water.  The light from the microscope may also waken them after a minute or two.
- If you are not ready to use the larvae immediately, they can be stored for several weeks in the water that was used to incubate them, so long as they are kept in the dark, or at least out of direct sunlight.
- If you need to add extra water, filtered or dechlorinated tap water or bottled water may be preferable to distilled water for this purpose. One very experienced incubator has found that rinsing or storing larvae in distilled water has killed some of hers,  whereas store-bought spring water has not.
- I don’t spend much time rinsing when they’re for my own use. But if the water is really dirty and I want to clean it up a bit, I dilute it with purchased spring water, although I never completely change out the incubation solution... I never rinse with distilled water as I find larvae don’t survive long in it... But I have successfully inoculated with 4 month old larvae when using store-bought spring water. (Edited from several posts in this thread.)
- When reverse osmosis water is used to store larvae, the osmotic pressure may be too low and put undue pressure on the larvae's excretory system. 
- It has been suggested that a greater weight of water above the larvae in their storage vessel may shorten their lifespan, so it may be best to limit the volume of water to just enough to cover the larvae - perhaps just 3 mm. 
- One user of this method has found that, if home-grown larvae are stored at between 60°F (15°C) and 65°F (18°C), some can survive for as long as 4 months,  and occasional fluctuations between 54°F (12°C) and 70°F (21°C) are unlikely to be a problem.  However, out of one group of eight 4-month-old larvae, only one successfully penetrated the skin,  therefore 8-10 weeks might be a more realistic expectation for larva survival. Also see, Hookworm larvae storage and survival.
- The larvae do need oxygen, which can become depleted at higher temperatures and by bacterial activity in the jar, although this is less likely to be a problem when using jars (e.g., 4 oz Specimen Cup/Container with Lid) rather than test tubes, and it can be prevented by opening the lid every few days and gently shaking the jar’s contents. 
- Even when kept in perfect storage conditions, the larvae will inevitably become progressively weaker as a result of the depletion of their fat stores.
Have a bandage ready that seals on 4 sides, or use a gauze pad with tape. Test bandages first to rule out allergic reactions.
Use a pipette to gather larvae in a few drops of water on the slide, but I don’t do this while looking through the microscope. Once the liquid is on the slide, I put this under the microscope and count the larvae, then add or remove liquid as necessary and recount. If there are a lot of larvae, I suck the water up until I reduce the numbers. Then, when I get the amount I want, I let the water from the slide drip on to a prepared bandage, or gauze pad with tape, and finally wipe the slide onto the gauze to get every last drop. Then I immediately apply this to my skin. I repeat this until I get the number of larvae I want, but can often get the whole dose on a single gauze.
I recommend 1-3 larvae at a time, depending on your goal dose. (See Hookworm dosing and response.) Place the bandage on a hairless, convex area of skin, such as your (shaved) leg. I don’t recommend the upper arm because the skin here is thin, and entry spots hard to observe, but a variety of sites have been used successfully. (See Body sites used for hookworm inoculation.)
Tingling should start after 10-20 minutes as larvae enter the skin, but let the bandage sit for at least 3 hours. (For more on this, see Inoculation with NA.)
Clean up supplies by freezing them in a ziplock bag overnight before disposal.
After a few hours or days, you should be able to observe the entry spots with the naked eye or a magnifying glass. If I didn't get a tingle and rash I would suspect failure.
Freezing is one of the easiest ways to kill off any remaining larvae at the end of incubation. Simply throw your slides, beakers, gloves etc. into a plastic bag and put them all into the freezer overnight.
This paper describes other methods you can use in your lab to kill hookworm larvae.
- Techniques to kill infective larvae of human hookworm Necator americanus in the laboratory and a new material safety data sheet
Note that hot water alone is not effective unless close to boiling point.
- Research safety extensively before starting (including reading The laboratory and safe practice), and have a doctor supervise you.
- Have a script for a course of an anthelmintic filled and ready before you start experimenting. Albendazole is more effective against hookworms than mebendazole, but isn't available everywhere. (See Terminating a helminth infection.)
- If using a donor, get them tested for HIV, Hepatitis B and C as a minimum. Strongyloides blood test is also recommended especially if you are on immune suppressing drugs.
- If using donor stool, take a peek at this under the microscope to see what’s in it when you first get it. Do a fecal float and check for eggs. (See Stool testing (egg counting).) If you see larvae in less than 24 hours, do not use, since it may contain Strongyloides.
- Get bloodwork done on yourself before inoculating (CBC, Hiv, Hep A+B) and afterwards (Hiv, Hep A+B, Stronglyoides).
- Do not prepare or open the jars of feces outdoors. Bathroom is recommended, as flies can lay eggs in seconds and flies will ruin the feces. Feces are their favorite food.
- If you put your container outside to incubate, make sure the temperature stays between 77ºF (25ºC) and 95ºF (35ºC).
Tips and adaptations
Several users of this method have made modifications, and suggested tips, as follows.
Using garden soil instead of vermiculite
Sterilised soil can be used on its own as the incubation medium, as described by Alana in her incubation protocol.
Sterilisation of the soil is necessry in order to avoid the possibility of contamination by cat or dog helminths such as Toxocara cati and Toxocara canis. There is also a possibility of contamination by larvae of free living nematodes which, although not harmful, can interfere with the counting of larvae for inclusion in doses of NA. One method of sterilising soil is explained by Alana here.
Someone else who is having continued success using sterilised soil has reported as follows.
Using more feces
Using only feces
One hookworm grower uses no soil, vermiculite or charcoal at all, and just adds enough water to produce a suitable consistency for the sample to stick to the slide without slipping off. He says there’s no smell and he gets lots of larvae. 
Using less water in the jar
This makes it easier to find the larvae.
Mixing feces to a thinner consistency
If the poop mixture slides down whatever it's placed on and ends up in the water at the bottom of the container, try a wider container to allow the samples to sit at a flatter angle.  Also try leaving whatever the poop mixture is placed on completely flat for 30 minutes before putting it in the jar. This helps to firm up the mixture and prevent slippage. 
Note that charcoal can rapidly soak up water, possibly making a mixture too dry, unless it's been dampened first.
Keeping the sample moist
NA larvae die in a matter of minutes if they are allowed to become dry. 
The sample will remain moist if the container holding it is covered by a solid lid. However, if it is covered with a coffee filter or cloth, daily misting may be necessary to keep it moist.
If you do mist the sample to keep it moist, only use a small amount of water. A garden or domestic sprayer may deliver too much.
One experienced hookworm grower likes to maintain his cultures at a humidity of 70-80%. 
Re-wetting the sample
After failing to find any larvae at all, in spite of repeated, careful examination of the water in her incubations jars, one hookworm grower decided to try re-wetting the fecal sample. She used the rest of the sample water from the bottom of the jar to irrigate the fecal blob on the slide. This resulted in the water "swarming" with active larvae, going from no larvae at all to many more than she could use. 
Charcoal - a possible cause of failure
Three growers have attributed failed, or reduced, yields to added charcoal.
But others have had no problems with charcoal, so the issue reported above may have been due to specific brands of charcoal, or to coincidence.
Someone else who reports having no issues with charcoal is using the type of charcoal found in water purifiers, and grinding this in a blender. 
Type of container may affect outcome
Using test tubes as containers
One incubator prefers to use test tubes because the much smaller quantity of water in a tube prevents the need for the extra step of leaving the larvae to settle to the bottom of a separate conical-shaped container.
The following incubation method using centrifuge tubes was employed by one team of researchers.
The faeces sample was first homogenised by stirring, then emulsified with activated charcoal and sterile water until the consistency resembled bitumen or tar. Approximately 7 g of this material was then smeared on to the upper third of 9 × 9 cm squares of 80 gsm copy paper and rolled into cylinders, culture material innermost. These cinders were then placed within 50 ml sterile centrifuge tubes, each containing 5 ml sterile water. (The use of copy paper produces a cleaner harvest because it degrades more slowly than filter paper.) The base of the paper rolls rested in the water, with the culture material near the top of the tube with the lid loosely secured. The tubes were then stored in a tube rack placed within a plastic box, which was lined with adsorbent paper soaked in water to maintain a humid environment. Finally, the boxes were placed in an incubator at 77°F (25°C) for 7 days. To harvest the larvae, the roll of paper was carefully removed and discarded from each tube, leaving the water containing the larvae at the base of the tube. 
Substituting a coffee filter for the glass slide
One incubator progressed from glass slides to coffee filter papers for holding the fecal samples. After placing each sample into the centre of a filter paper, she put this in the bottom of a jar, on top of a couple of canning rings from Mason jars. The rings allow the filter paper to touch a small quantity of water in the base of the jar, while preventing the paper from collapsing completely into the water. More details here and here. Eventually she abandoned the use of canning rings and switched to using 2 filters together, which helps to keep them upright. 
The ideal temperature for incubation is said to be between 73°F (23°C) and 86°F (30°C), but there are growers having success with a wide variety of temperatures.
- 90°F (32°C)
- This may be close to the upper limit. One very experienced incubator has reported that she has completely failed to get any larvae at all when the temperature has even temporarily approached 95°F (35°C). .
- 80°F (27°C)
- Keep mine at 80 and it works fine. 
- 79°F (26°C)
- 26.7 C works perfectly for me, harvest at 7 days. 
- We have purchased a "lab" incubator which we set to 26 degrees Celsius and results are amazing! 
- 72-75°F (22-24°C)
- If it drops lower or goes higher it gets wonky for me. 
- 68-72°F (20-22°C)
- Room temperature 20-22 degree celcius is ideal for me. I don't use an incubator. 
- 65-70°F (18-21°C)
- I keep them in a dark cabinet in a space that is between 65-70 depending on the season. 
- 62-65°F (17-18°C)
- Takes 12 days or so. (Without a heat source, just in a dark box in a basement.)
- 55-80°F (13-27°C)
- I incubate successfully at temps from 55-80 F. They begin to die at 50F on the low end and 90+F on the high end. 
- 50-70°F (10-21°C)
- My incubations get as low as 59°F and do fine. 
- I have found that cooler and even temps, given a bit more time, produce weeks of long-lasting larvae. (Without an incubator, just in jars in a towel-lined box in a stable part of the house.) When I started with rapid and warm I frequently killed them. 
Larvae incubated at higher temperatures may be ready after only 4-5 days. Those grown at lower temperatures might not be ready until 12-14 days, or later, but they will survive for longer.
Fluctuations in temperature should not be a problem so long as the upper and lower extremes fall within the range above.
Controlling the incubation temperature
Isolating single larvae
Using multiple slides.
Using a DIY micro-pipette.
Picking up individual larvae is much easier using a pipette with a micro-tip, and these can be created by super-gluing a "micro-tip glue applicator" nozzle to any standard pipette. These nozzles are available cheaply from Amazon and eBay: just search for "micro tip glue applicator".
Once you’ve glued a micro-tip to your regular pipette, it becomes very easy to pick up single NA larvae - or even individual TSO/TTO eggs - from a glass slide. And, to increase precision, just pinch the pipette’s tube, rather than squeezing the bulb at the end. 
Transferring larvae from slide to bandage
Hookworm larvae do not need to be cleaned if they have been grown at home for use by the same individual who provided the stool sample for their incubation. However, if that individual wishes to share the larvae they have grown, these can be cleaned using the following method.
Add one drop of commercially available 5% Lugol’s iodine to 5 ml of water (distilled, filtered or dechlorinated tap water, or bottled water). Then add 1 ml of this 0.02% iodine solution to 1 ml of water containing the larvae. After the larvae have spent 20 minutes in this solution, the iodine should be neutralised by adding a pinch of ascorbic acid (vitamin C) powder to the water. Neutralisation of the iodine is confirmed when sufficient ascorbic acid has been sprinkled into the water to make this completely clear.
If the cleaned larvae are not going to be used immediately, they should be removed from the iodine/ascorbic acid solution and rinsed in fresh filtered or dechlorinated tap water, or bottled water, and then stored in a small quantity of one of these same types of water. While distilled water is suitable for incubation and for briefly cleaning larvae, it appears not to be best for storing them, unless it is the same water they were grown in. (By the time the eggs hatch, distilled water that was used for incubation will have been modified by solutes from the feces sample.)
Since larvae that have been cleaned by any method will have a shorter shelf life than those that have not been cleaned, they are best left untreated until as close to the point of inoculation a possible. 
Also see this support group discussion about this topic.
Possible reasons for failure
Novice hookworm growers may need a period of apprenticeship before succeeding at the art of hookworm incubation, even when using this simple method. For more details, see Some find incubation surprisingly difficult.
Hookworm incubations can fail for a variety of reasons other than a lack of experience. Long-term hookworm growers, and even hookworm providers, can experience fallow periods when their incubations produce no larvae. One reason for this can be the egg donor’s diet (for more about this, see Promotion of egg viability by dietary manipulation) and certain brands of probiotic have been implicated by one grower, so diet should be considered if there is a continuing failure to grow larvae.
Suggestions / observations
If anyone using this method has any questions, suggestions for improving this page, or any other observations, please post these to the Helminth incubation group.
Other NA incubation methods
- NA incubation: very simple petri dish method by Steven (The second most popular method.)
- NA incubation: very detailed method by Alana. (This method provides information about every aspect of hookworm incubation.)