Given the incredible improvement in my health that resulted from controlled dosing with the hookworm, Necator americanus, I had to find some way to maintain my hookworm colony. However, in view of the fact that they’re not yet available from mainstream medicine, that two of the first four suppliers to offer hookworm commercially were closed down, and that another one was forced to relocate, I wasn’t willing to risk my future health to chance, so I decided to become self-sufficient. I therefore had to find out how to safely and successfully culture infectious hookworm larvae. After reviewing the literature, and performing a number of experiments, I developed the protocol that I now use. This is set out below for those who are interested, but anyone reading what follows must be sure to first read the next section, ‘Warning and Disclaimer’, because there are safety and, in at least one country, legal implications to incubating hookworm.
Warning and Disclaimer
- There is considerable risk involved in incubating hookworms, due to the highly infectious nature of their larvae. After successfully culturing hookworm larvae, it would be very easy to accidentally inoculate oneself, or someone else, with many hundreds of these organisms, with potentially very serious consequences for the individual’s health and perhaps also the future of helminthic therapy if news of such an accident were to reach the media.
- It may be illegal to breed hookworm in some jurisdictions, including the U.S., where they are currently classified by the Food and Drug Administration as biological agents (i.e. drugs), as defined in Section 351 of the Public Health Service Act and subject to an Import Alert.
- The information presented on this page is not advice, but for general information only. It has not been approved or evaluated by any governmental organisation concerned with the regulation of healthcare or drugs anywhere in the world. The accuracy, validity, effectiveness, completeness or usefulness of this information cannot be guaranteed.
- While the information presented here is related to the practice of the experimental treatment known as helminthic therapy, it is not intended to provide medical advice, diagnosis or treatment. The reader is hereby advised to always consult with a physician or other professional health-care provider regarding any health care problem or issue they might have.
- Anyone who chooses to make use of the information on this page does so at their own risk and no responsibility or liability whatsoever is accepted by the author or site editors for the use or misuse by others of any of the information on this page.
Purpose of incubation
My intention was to incubate the eggs produced by the colony of hookworms (N. americanus) that I already host in order to re-infect myself for the purpose of maintaining the beneficial effects that I had obtained from hosting these organisms.
My existing hookworms were purchased as larvae from a recognised provider of helminthic therapy, and I would never use hookworm larvae, or eggs, from any other source because I want to be certain that any organisms I introduce into myself are either harvested from my own body or from a donor who I can be confident is free from pathogenic organisms and, in the latter case, that the organisms supplied have been scrupulously cleaned.
The laboratory and safe practice
- I have created a designated work area where no food or drink is prepared or allowed.
- I always wear long sleeves, long pants, socks and shoes plus a lab apron while following this protocol.
- I always wear gloves of latex or a synthetic alternative, and avoid touching my skin (e.g., face) while working.
- I use a sharps container for broken sharp objects - usually an old pill bottle.
- I use a dedicated small, lined trash container with a lid, and empty this frequently.
- I disinfect all materials using two containers. First, I immerse the equipment into either undiluted 5-6% bleach or 2-3% ammonia[note 1]. Then, after about 5 minutes, I clean them in soapy water, before finally rinsing and drying them. (Guideline for Disinfection and Sterilization in Healthcare Facilities, 2008)
- I disinfect all work surfaces when finished and, to be confident that nothing remains alive, I use either undiluted 5-6% bleach or undiluted 2-3% ammonia[note 1]. I only ever buy and keep available one of these chemicals at a time and I store whichever one I’m using in a 0.5 litre (16 fl oz) container and apply this to the relevant surfaces, as required, using a sponge. I then wait at least 2 minutes in order to be confident that the larvae are dead. Once the clean-up is done, I thoroughly rinse the sponges about 10 times afterwards, to prevent the residue bleach or ammonia from rapidly destroying them.
- I absolutely never mix or work with bleach and ammonia together, since to do so would produce explosive and toxic gasses.
- Whenever I use bleach or ammonia, I make sure that there is good cross-ventilation to prevent the build-up of fumes.
Hookworms normally incubate in a faeces/soil mixture, so these are what I use[note 2], covered by granulated cork[note 3]. (The Harada-Mori technique Practical Guide to Diagnostic Parasitology didn’t work at all for me.)
The ideal soil is sandy loam (which ensures good drainage and oxygen supply), with a clay content of less than 15%, and preferably around 10%. (Pure sand and clay are unsuitable. Sand loses its moisture too quickly, while clay retains surface water, and also, if clay dries out, larvae can be prevented from moving through it.)
It might also help if it has a lot of organic material in it, perhaps 20%. However, I believe that any good loam-based potting mixture should work for the purpose I have in mind.
The soil should be close to a neutral pH of 7 and must definitely not be highly acidic. Since most plants require a pH of between 6.0 and 7.0., (link) most bagged potting mixes will be close to neutral (7.0) or slightly more acidic. However, the pH of some commercially available potting soils may also be as low as between 4.0 and 5.5., (link) especially if they have been formulated to meet the requirements of specific plants, such as camellias and azaleas, which are acid lovers. (Link)
I checked the pH of the soil I bought using a pH meter. It had a pH of about 6.3, so it required the addition of powdered lime, as it turned out, in the proportion of 1 part lime to 1 part soil, to produce a more neutral pH of around 6.9.[note 4]
Adding stool to the soil obviously renders it non-sterile, but in order to guarantee that everything that was alive in the soil and lime when I first brought them home was now dead, I baked them, separately, in an oven, for at least 30 minutes at a temperature of at least 82°C (180°F), after first having made sure that the soil was quite moist, but leaving the lime dry[note 5]. The resulting mixed soil/lime combination was less wet, which made it lighter when stored, less likely to smell, and less likely to have things eventually grow in it (such as mushrooms). Of course, it wasn't sterile once it cooled down, since it was exposed to the air, etc.
I used a large turkey baking container to hold the soil[note 6], covered over on top with aluminium foil, and a meat thermometer to measure its temperature during cooking, inserting the thermometer probe in several different places, and rotating the container to ensure even heating.
After each series of temperature checks that I made before the temperature reached 82°C (180°F), I cleaned the thermometer to remove any of the as yet non-sterilised soil.[note 7]
Since I wanted an even greater safety margin, I left the soil in the oven after 30 minutes baking and let it cool down slowly over several hours. Unlike food, I don't suppose it is possible to over-bake soil, as long as it doesn’t dry out.
After baking the lime in the same fashion, I also baked the covered granulated cork for over 30 minutes (although this might not have been necessary, since it was likely to be far cleaner than the soil) in the same container, after cleaning this in order to keep residue soil and lime from contaminating the cork. I didn't moisten the cork, but left it completely dry. Once the final soil and lime mixture had cooled, I stored it in 20 litre (5 gallon) garden buckets
Taking a plastic container (e.g., a plastic food container, tomato plant starter, or similar, approximately pint-sized [0.5 litre /16 fl oz], with holes in the base for drainage), I spoon some soil from one of my buckets into the bottom of the container to a depth of about 1.25 cm (½ inch), then, using a hand sprayer filled with dechlorinated water[note 8], I wet the soil sufficiently for the water to drip through after a few minutes. I then set the container on a suitable dish to catch the overflow water.
I deposit one bowel movement’s worth of stool[note 9] into a different container, then transfer this stool to the container that holds the soil[note 10]. I then add another 1.25 cm (½ inch) layer of soil to cover this[note 11]. I also make the top of this second layer of soil smooth in order to make it easier to scrape off the cork later. I then wet this second layer of soil liberally until water drips from the base of the container.
Next, I put a sheet of latex-free (I don’t know if this matters) gauze, cut to fit the container, on top of the soil. (When doing this, it helps to make a paper template that fits the container’s mouth, and then to cut the gauze to fit the template.) This gauze separates the soil and cork so that, when the cork is later spooned off, it won’t have soil contamination in it. I also use a paper towel to wipe the upper inside exposed sides of the container to remove any soil clinging to them.
Finally, I add a 1.25 cm (½ inch) layer of wet granulated cork on the top. (As cork tends to be rather hydrophobic, I pre-mix a sufficient quantity with dechlorinated water in a separate container ahead of preparing the culture sample.) I also use a different spoon to ladle it out of its bucket and mix it, since I don’t want to contaminate either the cork in the bucket or the newly wetted cork with soil.
Depending on how many larvae I intend to harvest, I might repeat this procedure, filling additional containers when I have subsequent bowel movements over the next few days.
In order to keep track of when a culture will be ready for harvesting, I write its starting and harvesting dates on my calendar. I also write both dates on an index card, which I place in or on the dish in which the culture container sits. The card may get soaked, but this is of no consequence.
For the next 8 days I keep the soil shaded and constantly moist, ensuring that the shallow pool of water that the container is sitting in never dries up. How much watering this requires depends on the ambient heat and humidity. In my situation, I have found that spraying the soil three times a day, for a total of around 40 sprays (15 - 10 - 15), is generally sufficient. Since the water that pools in the container contains filtered essence of faeces, it becomes brown over time and, if the conditions are right, it can start to smell. If this happens, I empty the pooled liquid daily and replace it with fresh dechlorinated water. This is in addition to the daily sprayings.
Developing hookworm larvae are reportedly unable to survive temperatures below 10°C (50°F) so the temperature should preferably be kept over 18°C (64°F), and definitely below 35°C (95°F). Although the ideal temperature is reported to be between 27°C (80°F) and 32°C (90°F), I have had excellent results with temperatures in the range of 21°C (70°F) to 27°C (80°F). I have, however, completely failed to get any larvae from cultures when the temperature has approached 35°C (95°F), even temporarily.
- Page 30, Integrated Guide to Sanitary Parasitology
If I lived in a cooler climate, which necessitated warming the containers, I might try using a low wattage light bulb or an aquarium reptile heat rock. If placed inside an appropriate container, either of these might provide a suitable temperature increase of, say, 3°C (5°F) to 6°C (10°F). The temperature could be controlled using a temperature switch, e.g. the Red 12V Heat Cool Thermostat Temperature High Low Alarm Control Switch -55-120C. If connected to a 5.0V DC power supply (e.g., an old cell phone charge adapter), this might provide a good low-cost DIY incubator.
Hookworm eggs hatch in about 1-2 days and, by days 5 to 7, the 2nd stage larvae have become 3rd stage larvae and will migrate upwards to the top of the soil.
Hookworm larvae cannot swim and, if put into open water, will sink to the bottom. However they can climb against gravity by using surface tension. So, when the 3rd stage larvae develop, they use this modus to crawl up into the grains of cork on top of the soil as they search for the surface.
By day 8, the larvae should be ready to harvest.
In their natural environment, hookworm larvae crawl up blades of grass with the morning dew - hence the name ‘dew itch’, (, ) so, about 2 hours before I remove the cork to harvest the larvae, I spray the culture thoroughly to make sure they are able to get as high as possible in the cork, although I don't know how much difference, if any, this makes, or what the ideal timing would be.
For the previous 3 days, and from this point onwards, I’m particularly careful to always wear barrier gloves, a long sleeved shirt and pants, shoes and socks, plus a lab apron when directly handling any of this material.
I half fill a silicone snow cone cup with dechlorinated water. Then I use a spoon to scoop off some of the cork from the top of the culture and place this into the water in the cone, thoroughly stirring the cork into the water. Once there is sufficient cork in the mixture, I add some more dechlorinated water to bring the surface of the mixture to about 0.6 cm (¼ inch) below the rim of the cone. Finally, I set the cone aside and wait approximately 12 hours for the larvae to settle to the bottom. The cork floats immediately to the surface.
For the next step, I use a micropipette/minipipette (MP) fitted with a tip that has been cut off at the first division from the narrow end in order to widen the hole sufficiently to prevent it becoming blocked with debris that has settled to the bottom with the larvae[note 12]. I depress the MP's plunger before putting it into the water (doing this with the pipette under the water will produce bubbles that might disturb the sediment and distribute the larvae). Next, holding the plunger down, I insert the MP into the water and down to the bottom of the cone. Then I release the plunger and slowly raise it, with its tip full of liquid, out of the water, momentarily tapping the MP on the edge of the cone to dislodge any drops of water that may be clinging to its exterior, before transferring the drawn up liquid to the slide[note 13]. (Sometimes, it is also necessary to scrape back into the cone any bits of cork that have clung to the MP or to my glove, although I could avoid the need to do this by using a teaspoon to ladle off the cork before this part of the procedure.)
I then depress the MP's plunger to deposit the liquid from the tip onto a microscope slide. If everything has gone according to plan, there might now be a hundred or so hookworm larvae in that single drop of liquid.[note 14]
After putting the slide under the microscope, turning on the light, setting the total magnification at 40X, and adjusting the focus, I expect to see either motile larvae or resting ones that look like little cuticles or sticks. The latter usually become active after a few minutes or so, once I turn up the intensity of my microscope's halogen lamp.
If I don't find any larvae, I repeat the procedure with the same cone a number of times, and then repeat it with any additional cones that I may have prepared. Since the larvae can be very unevenly distributed in the cork it is possible for one cone to contain many larvae and another to contain few or none.
Storage and clean-up
If I want to store any larvae, I next use a transfer pipette to draw up liquid from the bottom of the cone and expel it into a Glad container. I repeat this, checking occasionally as I’m going along to see if I’m still getting larvae, until I’m no longer doing so, or until the container is about 75% full. I’ve punched about 8 small holes in the Glad container’s top, with one end of pointed scissors, to let air in, although I don’t know if this is necessary, nor how long the larvae could survive in a fully closed case.[note 15]
I store the filled Glad containers in an unplugged mini fridge (e.g., the Caldura 17 litre Compact Mini Fridge) on the assumption that, if my home were to burn down, my precious stock of larvae would have a better chance of survival if protected by the fridge's insulation. I also keep my important papers, including inoculation records, in the fridge. Since I typically open this fridge about once each day to retrieve some paper or other, this arrangement ensures that the air in the fridge circulates periodically, although the larvae must use very little air.
Stored larvae need to be protected from direct sunlight, and checked from time to time to ensure they don't dry out through evaporation. If necessary, I add a little distilled water rather than using more regular dechlorinated water which would gradually increase the concentration of minerals in the remaining water. I don’t know what concentration of solution the larvae can tolerate.
Although it is clearly best to use the larvae while they are still young and vigorous, they may, if stored in this way, at around 21°C (70°F), live as long as 15 weeks, whereas, at 35°C (95°F), they will be more active and, as they do not feed, will die of exhaustion in less than 3 weeks. This is inconsistent with the the findings of Udonsi & Atata, 1987 (Necator americanus: temperature, pH, light, and larval development, longevity, and desiccation tolerance), a fact that I currently can’t explain.
Since the material left over from this protocol contains infectious organisms, I don't simply throw any of it out with my domestic waste. First I freeze leftover stool, soil and cork for a couple of days, then let it thaw out and, finally, with gloves on, mix it thoroughly (break it apart) and add it into either undiluted 5-6% bleach or undiluted 2-3% ammonia. I make sure to add enough bleach or ammonia to keep the resulting solution at least 50% of its original concentration. I then wait at least 1 hour before draining off the excess bleach or ammonia using a dedicated strainer, and triple bagging the remaining solid materials in plastic bags, before putting them outside the house in a refuse sack along with the rest of my regular rubbish. As an alternative, I will sometimes simply boil all the materials for 10 minutes and then triple bag them after they have cooled down.
To ensure that I have clean slides the next time I need them, I thoroughly wash the ones that I have used with a clean cloth or sponge, film-free soap, and water, then rinse them several times[note 16]. Finally, I pick up the slides by their edges, replace them in my slide case and leave them to dry.
Counting the larvae
Since a single drop of water can contain hundreds of larvae, it is essential for me to use a microscope to count out the required number. The dose for an adult could be anywhere from 1 larva to as many as 50 larvae, with a first dose for an adult being either 3 or 5. For more about this, see the Hookworm dosing and response page.
I use a micropipette/minipipette (MP), with tip intact, to remove a drop of liquid from the bottom of the Glad container, then place this on a slide.
10 larvae per drop is about right for easy counting. If there are too many larvae in each drop of the sample to be easily counted, I might expel very small drops onto a slide and add dechlorinated water to each of these, or I might divide a drop on a microscope slide with the MP tip, and then add water to these daughter droplets. The larvae do tend to congregate together, but this isn't a problem.
At this point I wake the larvae with the microscope light to be sure they are still alive and will be awake when they are put onto my skin. There is the question of whether or not to count the larvae that don’t wake up and move on the slide (they look like sticks). I don't count them, and it works out about right, since when I count the bumps on my arm this closely corresponds to the number of active larvae I counted out.
To transfer the larvae from the slide to a contact lens case, in readiness for the inoculation, I MP up the drop and expel it into the clean case. Then I MP up fresh water from a shot glass and expel this onto the same spot on the slide, gently stir the drop with the tip of the MP, and then MP this up to transfer it to the case, sometimes repeating this several times. Doing this doesn’t seem to leave too many larvae behind. I keep track of the total number of larvae as I add further drops to the case, which will eventually contain, say, 20 larvae.
The fabric pad of the dressing will need to be in firm, direct contact with a patch of clean, hairless skin (to prevent ripping hairs out when I eventually pull the dressing off) in an area that is convex and not subject to much movement, so I use the inside of my upper arm.
In order to sterilise the area, I wipe some alcohol on it and wait briefly until this has completely evaporated, which usually takes about 45 seconds. (I’m careful not to clean or disinfect the area with anything that might leave a residue.) In the meantime, I lay out one large adhesive dressing so that its fabric pad is exposed but leaving the protective plastic backing still attached to the adhesive and folded back against the table. I only remove this backing as I am applying the dressing to my skin.
I use a transfer pipette to suck up the fluid from the contact lens case and transfer it to the dressing. This might require several transfers and, to ensure that any larvae left in the case do not dry up during this operation, I immediately deposit more fresh dechlorinated water into the case if it becomes empty.
In order to ensure that I have collected all the larvae, I squirt a little fresh water into the case then draw it back up and transfer it to the dressing, repeating this flushing action a number of times in different locations within the case.
If the dressing becomes saturated before I have completed this process and the water begins to bead up on top of the dressing, I put the case aside for a moment, after adding a little more fresh water to it, then draw up the excess water from the dressing and put it back into the contact lens case. I then attach the first dressing to my skin, stretching it slightly to ensure firm contact across the entire dressing.
If I haven’t been able to transfer all the liquid from the case, I quickly prepare a second dressing, and repeat the same process with this one. Once the second dressing is prepared, I apply this next to the first one on my arm. Then I leave the dressing(s) in place for at least a couple of hours.
When I feel an itch, I then swallow the maximum permitted dose of diphenhydramine.[note 17] This does not harm the larvae, but does make me feel drowsy, so I always inoculate myself at a time when I know I will be able to lie down for a number of hours, if necessary. In order to give the larvae plenty of time to pass through my skin, I obviously don’t use any Benadryl cream at this point, or apply a hair dryer (see below). However, I may vigorously slap the dressing to give me temporary relief if the itch becomes unpleasant before the Benadryl takes effect. I doubt that this will harm the larvae and it may in fact wake up any that are slow to action.
(N.B. Gradually, over the last six or seven years, my itching from the larvae has greatly reduced, and therefore currently I don’t need to use the liquid Benadryl anymore, although I still do need to use the creams and hair dryer later on.)
After a few hours, I remove the dressing and apply a combination of extra-strength (2%) diphenhydramine hydrochloride cream [note 18] with a 1% hydrocortisone cream, and an electric hair dryer. I find that applying heat to the site of entry - up to the point of momentary pain - effectively kills any remaining itch for a number of hours. (A Miraculous Cure for Bug Bite Itching) [note 19]
For the first 5 days after inoculating, I avoid clearing my throat and spitting out any phlegm that might be produced, so as not to spit out any larvae that might happen to be migrating from my wind pipe to my throat at this time. I actually find this rather difficult to remember, especially when running in the gym, so I wear a wrist band during this time period to remind me.
Maintaining my colony
After my initial dose of hookworms, I followed this up, 6 months later, with another similar dose. Since then, I have given myself approximately 20 larvae every 6 months to keep my colony young.[note 20] For convenience sake, I have considered doubling the dose to 40, and then dosing only once a year. The problem with this is that with a higher dose I would be far more likely to have side effects, such as cramping and diarrhea.
While only dosing every six months, I prepare a fresh culture monthly in order to always have fresh larvae available, in case I ever lose my resident colony. I also keep the last two months production before disposing of them.[note 21] If I were dosing yearly, and also only doing a yearly culture, this would obviate a lot of work, but would also obviously mean that I wouldn’t have any back-up in case I somehow lost my colony.
Everyone is different in how they respond to hookworms and research has suggested that, in some people, the presence of older worms may make colonisation by younger ones more difficult, so it would be impossible to know how many worms I’m carrying unless I had a pill cam examination. But, at a guess, and assuming that hookworms live on average for 5 years and have a 50% survival rate, my present schedule will maintain a colony of about 100 hookworms that won’t get old and die at the same time. For more about dosing, see the Hookworm dosing and response page.
The estimates for the number of hookworms that begin to create a risk of anemia for someone with normal iron in their diet and normal iron absorption vary widely. One opinion is that a person would need to have somewhere around 2,500 to 5,000 worms before anemia would become a risk, giving someone a safety multiple of approximately 25 to 50 times a normal colony's size. A contrasting position is that around 300 to 400 hookworms could create a risk of anemia. So although a person with a normal sized colony wouldn't be expected to develop anemia under either scenario, given the current uncertainty, anyone with hookworms probably would be prudent to monitor their iron levels. So long as iron levels are checked periodically and an iron supplement taken as and when required, there is no cause for concern about developing anemia - or any other nutritional deficiency - from hosting therapeutic numbers of hookworms. For more detail about this, see Helminthic therapy and nutritional deficiencies page.
I don't want to risk allowing my entire colony to become geriatric. Moreover, dwindling numbers could result in only one sex being left, in which case I wouldn't be able to reinfect myself as no eggs would be produced. For example, if I had only 4 worms, the probability of getting all males would be ½ x ½ x ½ x ½ = 1/16. I would also have a 1/16 probability of getting 4 females. So the combined odds of getting all of only one gender is 2/16 = 1/8. (If I had 10 worms the odds would be about 1/500.)
Another problem with having small numbers is that my colony could suffer reduced genetic diversity. With fewer breeding pairs, my population might be in a genetic bottleneck and become very inbred.
Promotion of egg viability by dietary manipulation
Having noticed that the production of larvae from my stool is variable, and having controlled for all other possibilities, I concluded that what I eat must be influencing the viability of the eggs produced by my hookworms. I therefore drew up a list of foods, herbs and spices that are claimed to be anthelmintic and began testing these individually.
The first four items that I selected each completely eliminated larva production. These were: regular (organic) English walnuts (not the black ones); carrots (the roots as well as the carrot tops); garlic and turmeric. In view of this, I decided that, until I was able to establish the safety of every other item on my list, I would totally avoid all of them for a week or two before beginning a new culture.
I then continued to experiment for several years and found that everything from fish oil to a 1-a-day multivitamin limits my ability to culture larvae. So, in order to obtain top-up doses, I have needed to eat a special diet of only hard boiled eggs, boiled chicken, salt, butter and water, and nothing else, for 30 days before starting a culture.
But then I discovered that taking prednisone for a few days would enable me to get a tiny culture of approximately 10-20 larvae. This was a wonderful development, and the first progress I’d made with this in several years, although the prednisone did make me very anxious/paranoid while I was taking it, possibly due to my multiple chemical sensitivity.
The next step will be to try combining prednisone with a shortened period on the special diet to see if there's a synergy that might allow me to successfully culture more larvae without the need for prolonged dieting.
Someone else who incubates hookworms has reported that a 9 day course of amoxicillin stopped egg production for at least 16 weeks, but that eggs were once more being produced at 20 weeks.
Another worm host who regularly checks the egg output of his worms has reported that they produced zero eggs for a time after twice drinking coconut milk. And yet another individual has found less eggs when he consumes 2 large dessert spoons of coconut oil daily, but normal amounts of eggs when he only eats 2 smaller dessert spoons of the oil each day. This confirms the ability of coconut to affect helminths in some people, as well as demonstrating the dose-dependent nature of this effect.
Yet another would-be hookworm incubator has reported that he had no success while taking a zinc supplement in doses he had increased slowly up to 130 mg per day to treat type 2 diabetes. Three weeks after he stopped all zinc intake, he found NA larvae in his incubations for the first time. 
A 2022 review paper found a number of foods that have shown some level of anthelmintic effect, including papaya, walnut, pomegranate, asparagus, squash, nutmeg and coriander.
The issue here is the potential effect of these items on larva production, not their possible adverse effect on the health benefits produced by the hookworms. This latter topic is addressed separately in the Human helminth care manual.
- Techniques to kill infective larvae
This paper (Techniques to kill infective larvae of human hookworm Necator americanus in the laboratory and a new Material Safety Data Sheet) along with a few experiments and consultations with others lead me to the conclusion that 90% isopropyl alcohol, 3% hydrogen peroxide, Lysol, 2% Glutaraldehyde, 10% Formalin, full strength Dettol, 2% Chlorhexidine, and 10% Povidone Iodine are all unsatisfactory for quickly killing 3rd stage hookworm larvae. Virtually pure ethanol (sort of) works, but is expensive and unavailable in many places. Very strong sodium hydroxide (soda lye) is very effective, but also extremely harsh to work with. Therefore, I currently have four practical off-the-shelf methods for killing hookworm larvae: Two slow methods: freezing, and boiling in water; and two relatively fast methods: undiluted 5-6% sodium hypochlorite bleach, and undiluted 2-3% ammonia.
In repeated tests that have been conducted with either undiluted 5-6% bleach or undiluted 2-3% ammonia the larvae stopped moving within 2 minutes. On this basis I'm assuming that, at this point, they are dead, or at least no longer infectious.
If the bleach or ammonia is mixed in with stool, soil, water or cork, it becomes diluted, necessitating that I leave it to work for considerably more time.
- Instead of soil, I could possibly have used charcoal (e.g., granulated activated carbon/charcoal) which I would have been able to find at aquarium supply shops, as well as online, e.g., Finest-Filters 1000g Granulated Activated Carbon / Charcoal for Aquarium and Pond Filters. Another option would have been vermiculite, which is available from garden centres and online, e.g., Vermiculite - natural incubation substrate - 5l Bag - 2-4mm Grain Size - Incubation Substrate. However, I would have had to grind both of these materials and sterilise them by baking them.
- The cork I use is granulated, 20–40 mesh size, and is claimed by the supplier to be in its natural state, with no added chemicals.
- The pH scale runs from 0 to 14, with 7 being neutral. So from 0 to 6.9 is the acidic range, and from 7.1 to 14 is the basic range. Since my soil was not too basic, I was not faced with the problem of finding a product to make it less basic, and thereby closer to the ideal neutral pH of 7.
In some places it has been reported that many of the products for making soil less basic seem to come in a very large grain size, which might have meant that I would either have had to hunt around to find a product that was in a sufficiently fine powdered form, or have been prepared to identify a way to grind what I did find into an acceptable powdered grain size, perhaps using an electric cheese grater, an electric pepper/spice mill, or an electric mortar and pestle, e.g., Glen Mills.
- Web sites warn that the soil might smell when baked, but this wasn’t a problem when I baked either the soil, the lime or the cork. I have also read warnings that birds can be extremely sensitive to indoor air pollutants, which could kill them, but this wasn’t an issue for me, since I don’t own any birds. (How to Beware of Deadly Indoor Air Pollutants That Can Harm Your Companion Bird) They also warn not to get the soil over 93°C (200°F), e.g., How to sterilise potting soil.
- In some places they sell inexpensive large aluminium foil turkey baking pans. If these had been available, and I had decided to use something like this, I suspect these would have been very difficult to get clean due to the numerous small crevices on their inner surfaces, so I might have needed to reline them on the inside with fresh aluminium foil before baking the cork in order to keep the residue soil and lime from contaminating the cork.
- To be sure that my thermometer was reasonably calibrated, I had previously tested it in ice water 0°C (32°F), as well as boiling water 100°C (212°F), and adjusted it to be the most accurate in its upper temperature range. (If I were doing this at an extreme altitude, I would have needed to take into account the fact that water boils at slightly different temperatures at different altitudes.)
- I use dechlorinated water for my cultures, although I’m not completely sure that this is necessary. Dechlorinated water is easily prepared by filling a shallow plastic container, (e.g., 4 litre [1 gallon] size, 30 cm x 20 cm x 10 cm [12” x 8½” x 4”]) with cold tap water, and then letting it stand with the lid off for 24 hours. Tiny bubbles form on its inside surfaces, which I remove by tapping the container’s sides, and then scraping the remaining ones off using a
kitchen knife. Then I snap on its sealing top, and use the water as needed. I don’t use distilled water, since I don’t know if the larvae can tolerate the osmotic problems that might be associated with this, and I'm guessing that normally mineralised water is what they are most comfortable in.
Many areas use chloramine instead of free chlorine to disinfect drinking water. If this were my situation, I would first carry out a test to establish whether this had any adverse effect on the larvae. If it did, then I would either have to investigate using one of the products designed for tropical fish owners, e.g. Seachem Prime, (this site presents a list of them); investigate using bottled water, distilled water, or water from a local natural source; or even, as a last resort, consider installing an expensive and complicated multistage filter system.
- Chlorine and Chloramine in the aquarium
- Chloramine Facts
- Anyone knows how to remove chloramine from drinking water?
- Frequently Asked Questions - Chloramines
- Chloramine - Wikipedia)
- The stool should not come into contact with either urine or chlorinated water. Since I don’t want any chlorinated spray getting in my cultures, I deposit the stool away from any sink, and also keep my cultures away from them. Obviously, I don’t retrieve my stool from the toilet. I find it easiest to deposit it directly into the container while in my bathroom.
- If my stool is soft, I use a dedicated knife to cut it up and mix it with the soil, adding first some soil, then several lumps of stool. Then I spray it down, add more soil, spray it down again, and so on. This creates a kind of heterogeneous construction aggregate which prevents the stool from forming an impenetrable plug in the culture container, causing water to pool on top of it instead of draining through the soil/stool mixture. With very hard stool this isn't necessary.
- One advantage of using soil is that it covers the stool, greatly reducing faecal odour problems.
- If a tip does become clogged I clear it by running water through it backwards.
- I do this to prevent a build-up of excess liquid on the slide, from where it might run over the sides of the slide and make a mess, getting onto the back of the slide and causing this to stick to the microscope stage, which would then need to be cleaned.
- Interestingly, my yield of larvae has dropped greatly over the last few months. Instead of a hundred larvae per drop, I have recently been getting around 6, with a total for the entire culture of only 150 to 200, rather than thousands. I‘m not entirely certain of the reason for this development, but it could be due to aging worms, unduly hard stools, or, as mentioned under “Promotion of egg viability by dietary manipulation”, it may be the result of eating foods, spices or herbs that are affecting eggs production. Given my most recent experimental results, mentioned at the end of the dietary manipulation section, I think this last possibility is overwhelmingly likely to be the main cause of the problem.
- If there was a need to further concentrate the larvae, I could empty the Glad containers containing the larvae into an empty cone, and also rinse the cases into that cone to ensure that all the larvae had been transferred. I would then wait about 36 hours before drawing off the surplus water from higher up the cone. Then I would draw up what was left at the bottom of the cone, before rinsing the cone several times, when empty (as I do with the contact lens cases when preparing a dressing for inoculation) and then I would store the rinse water in another Glad container.
- I use white Dove bar soap for the initial wash because it doesn’t seem to leave a film, but any film-free cleanser would work just as well. After the slides have been rinsed in regular water, I rinse them twice more in distilled water (Distilled Water - 5.5 Litres) in a bowl, swirling them around in it, changing the water, and then rinsing them again. During the whole of this process, I only hold or touch the slides by their edges.
In order to remove absolutely all of the mineral deposits, I often take another bowl, rinse it 4 times using a small amount of fresh distilled water for each rinse and swirling this around before disposing of it (thus hopefully removing any leftover mineral deposits from the bowl’s previous regular washing) and then refill the bowl with fresh distilled water. I don't wipe the bowl during this process because a towel might add contamination. Then I process the slides, one at a time, in assembly line fashion, shaking off any clinging water left from the first bowl, swishing each of them in the second bowl, then again shaking off any excess water, before putting them in the slide case.
If I lived in an area where the tap water was really hard and, as any minerals already on a slide may slightly contaminate the distilled water I'm using in any given bowl, I might need to repeat this procedure in a series involving multiple bowls and repeated distilled water rinses before finally putting the slides away. I must also carefully clean the mechanical stage of my microscope before each use, to prevent any dust, etc., from contaminating my clean slides. Even after all this, I’ve found that, over time, my slides have accumulated imperfections and marks that can’t be removed. My solution to this problem is simply to dispose of the imperfect slides and use new ones.
- I wait till I feel the itch so that I have a clear indication that the worms are alive and making a start on their journey before I take the oral diphenhydramine. If I had just checked the larvae under a microscope and established that they are all active, I might not need to wait for an itch and could take the medication about 15 minutes before inoculating.
- Diphenhydramine is one of the best antihistamines for helminth hosts to use, although it typically causes drowsiness. Other worm-safe antihistamines that generally cause less drowsiness are
- loratadine (Claritin)
- and fexofenadine (Allegra, Telfast).
- cetirizine (Zyrtec, Reactine)
- levocetrizine (Alcet, Allear, Curin, levcet, Seasonix, T-Day Syrup, Teczine, UVNIL, Vozet, Xaltec, Xozal, Xuzal, Xusal, Xyzal, Zilola, Zyxem)
- desloratadine (NeoClarityn, Claramax, Clarinex, Larinex, Aerius, Dazit, Azomyr, Deselex and Delot)
- and possibly acrivastine (Semprex-D in the US), all of which may potentially harm helminths.
- Diphenhydramine has local anaesthetic as well as antihistaminic properties, which makes it doubly beneficial for treating a hookworm inoculation rash. Topical products containing 2% diphenhydramine hydrochloride are available in several forms, e.g., Benadryl Extra Strength Itch Stopping Cream, Benadryl Extra Strength Itch Stopping Gel, Benadryl Extra Strength Spray and Benadryl Extra Strength Itch Relief Stick. These, and similar products, are available in both the US and UK - from Amazon, eBay and other outlets - and may also be available in other countries.
- In addition to the methods I use to deal with the inoculation itch, there are details of approaches used by others on the Hookworm inoculation rash page.
- I make a record of all my inoculations to help me keep track of my likely colony size, and to help me determine when I need to reinoculate. I also mark the dates of inoculations on my calendar.
- I record the date of each harvest on an index card, which I cut to a short, narrow strip to fit the side of the Glad container, then tape this in place.
I’ve found that the sort of materials listed below are currently adequate for my culturing of hookworm, although the suppliers mentioned here are not necessarily the ones I used, merely sources of similar items.
This list, as well as any items mentioned or cited previously, is offered solely for educational purposes and its inclusion is not intended to be encouragement to anyone to emulate my practice of hookworm culture. The Warning and Disclaimer printed above apply.
Common sense should be used when interpreting this list, because I might have been able to find substitutes for the items listed below that would have performed the same functions. For example, when dimensions of items are given this usually doesn’t mean that I would have had to use exactly those dimensions. Such numbers are included only so the reader has a rough idea of what I’m describing.
I have included graphics of some of the listed items for reference if and when the links expire.
- Micropipette/minipipette, 100 microlitre e.g., Fixed-Volume 100 µL (±0.3 µL) MiniPipet (see graphic 3A)
- Micropipette/minipipette tips (see graphic 3B)
When I bought my mini pipette tips I had to check carefully to make sure that they were compatible with my particular mini pipette, e.g.
- Transfer pipettes (see graphic 3C) e.g.
- PH meter e.g., Mosser Lee Soil Master PH Meter (see graphic 3D).
- Funnel for filling water sprayer bottles e.g., 90mm Plastic Transparent Funnel for Kitchen (see graphic 3E).
- Potting soil e.g., Scotts Miracle Grow Potting Mix Comp 8lt (see graphic 4A)
- Powdered limestone
I found this necessary for making my soil less acidic, e.g.
- Granulated cork (see graphic 4B)
20 – 40 mesh size, 1 to 3 kg (2 to 5 lb) (One 20 litre [5 gallon] garden bucket will hold about 1 kg [2⅓ lb] of cork.)
I was able to find a local retail cork supplier but, if this hadn’t been possible I would have investigated these online sources to ascertain whether they would be able to supply cork in small amounts that was free of any added chemicals, e.g.
I investigated the possibility of grinding my own 20/40 mesh granulated cork, which looks entirely possible, and not at all technically demanding, though I won't be pursing this because it is far easier, and probably far cheaper, to buy it already in granulated form. If I were to grind my own cork, I would need to buy an electric cheese grater (using the fine/small grating material), two sheets of filter mesh (one 20 mesh, the other 40 mesh) and then find a cheap supply of raw natural cork scraps, or use widely available store-bought natural cork bottle corks, although I suspect the latter option might be quite costly.
Possible sources of 20 & 40 mesh screen:
- Electric cheese graters e.g.
- Raw cork
This should be obtainable from the sources listed for 'granulated cork' (see above)
- Trash container (lid type)
e.g., Brabantia Pedal Bin with Plastic Bucket, 3L - Brilliant Steel (see graphic 5A)
- Plastic containers - 0.5 litre (16 fl oz) - for holding bleach or ammonia.
- Plastic containers - 0.5 litre (16 fl oz) - to hold the cultures.
A small supply of approximately pint-sized containers (to hold the cultures) with holes in their bases for drainage. Plastic tomato plant starters, or similar, would have been suitable, but I made my own by drilling holes in the bottoms of plastic food containers.
I am very likely being too pernickety but, for all the equipment that I use to hold liquids or that will come into contact with moisture, I avoid using plastics that might contain Bisphenol A (see Why All the Fuss About Bisphenol A (BPA)) so I stick with plastic types 1, 2, 4, and 5. I also tried to keep to these types of plastic for other items that I used for this protocol - at least when I could tell what the plastic type was.
- Plastic containers - assorted; to hold miscellaneous items.
- e.g., 2 litre (64 oz), 15 cm x 20 cm x 9 cm (6” x 8” x 3½”) and 2 x 15 litre (16 quart) containers
- e.g., 15 Litre Plastic Storage Box Container With Clip On Lid and Handle (see graphic 5B).
- Contact lens cases - I use three of these, e.g.
- Spray bottle (for dechlorinated water) e.g.
- Used plastic pill bottles - miscellaneous, for sharps, etc.
- Shot glasses, 2 – 4, either plastic or glass e.g.
- Reusable silicone snow cone cups with wire cup holders, 2 packs of (4 total) e.g.
- Back to Basics SIT10895 Snow Cone Cups and Holders, 2-Pack (see graphic 5H)
- Instead of snow cone cups, a well-equipped lab would probably use something like an Imhoff cone, e.g., Sedimentation Cone to Imhoff graduation up to 1000ml or even a separatory funnel.
- I could also have used something like this Gold Martini Glass (see graphic 5I).
- 20 litre (5 gallon) buckets, with lids
How many buckets I was going to need obviously depended on how much cork and soil I was going to store. One bucket will hold about 1 kg (2⅓ lb) of cork, or about 25 kg (50 lb) of soil and lime mix.
- Bin organisers - for small items
- Mine look something like these: Faithfull Plastic Storage Bins with Wall Mounting Rails (12 Pieces) (see graphic 5J).
- Alternatively, any of these would also work well:
- Mini fridge e.g.
- Caldura 17 litre Compact Mini Fridge (see graphic 5K)
- Plastic washing up bowls
I have used three of these in the past (approximately 10 cm deep x 30 cm x 30 cm [4” deep x 12” x 12”]) and they can still come in handy, but I now generally use the sink for all washing purposes. e.g.
- Dishrack - I find it useful to have a separate one for drying washed lab equipment.
- Lehigh Spontex 17005 Bluettes Knit Rubber Glove (see graphic 5M).
- Disposable exam gloves in latex, nitrile or vinyl
It’s far cheaper to reuse the gloves a number of times, but if these are put back on too soon, they may still be damp from the previous use and hard to get back on. In this case, I use a hair dryer to dry them out. Alternatively, I may place the dryer on my wrist, while it’s set on cool rather than hot, and use it to inflate the glove (which at that moment is stretched over both my wrist and the dryer’s mouth) as I work the glove back on. Yet another option is to rotate several pairs.
These are widely available at local pharmacies and online, e.g.
- Vinyl lab apron e.g.
- Black apron water proof resist vinyl back chef cook butchers pocket bib halter
- Lab Apron No. 2, heavy vinyl
- Hamilton Bell Co. Inc., No. 5250, 27” x 42” (see graphic 5O)
I found mine at an online science supply site. Some college bookstores may also carry these for their chemistry students. On mine, the apron’s tie cords were made of plastic, and quickly broke, so I replaced them with used shoe laces attached to the apron with shoe goo and staples.
- Shoe goo (see graphic 5P)
- Kitchen sponges, several e.g. 3M O-Cel-O Handy Sponge Power Pack 7274T, 4-Count (see graphic 5Q). I really like this type of sponge for cleaning up, and I also use them to make a bed to rest my hydrometer on when I’m incubating whipworm eggs.
- Terry cloth towels - 2 or 3, small
- Paper towels, 1 roll
- Plastic refuse bags - for triple bagging and disposing of old soil and stool
- Aluminium foil, 1 roll
- Turkey roasting pan, aluminium foil type, approximately 40cm x 33 cm x 8 cm (16”x 13”x 3”) e.g.
- Bleach or ammonia - (Either) bleach, by the litre, (or gallon), 5-6% sodium hypochlorite (Or) ammonia, by the litre (or gallon), 2-3%.
- Alcohol, one bottle 90%+ (If I had been using alcohol wipes, I would have needed to be sure they didn’t leave any residue.)
- Adhesive dressings, one box
- Diphenhydramine - A liquid form of this drug is preferable to ensure more rapid absorption, e.g.
- Diphenhydramine Hydrochloride cream, maximum strength (2%)
This is only occasionally available from Amazon.co.uk, but can be ordered from Amazon.com (US), e.g.
- Dr. Sheffields Anti-itch Cream with Histamine Blocker - 1.25 Oz
- Benadryl Itch Stopping Cream, Extra strength-1oz. I always make sure that I have plenty of this cream in stock.
- 1% hydrocortisone cream (This hasn’t always been necessary for me.) e.g.
- Electric hair dryer
- e.g. Eti Hair Dryer (see graphic 5S). Using a dryer hasn’t always been necessary for me.
Helminth incubation discussion groups
If anyone using the information on this page has any suggestions for its improvement, or any other observations about it, please post these to any of the following groups.
They will then be collected and added to this page.
Other NA incubation methods
Some of these hookworm incubation protocols are greatly simplified but, even if they seem superior to the protocol presented here, it might still be useful to first read the whole of this page to provide a full understanding of all the issues surrounding the practice of hookworm incubation.
- NA incubation: very simple Harada-Mori method by Sarah (The method most used by home growers and featuring many supplementary details gleaned from user feedback that will be useful to anyone using a different method.)
- NA incubation: very simple petri dish method by Steven (The second most popular method.)