This simple method will separate Necator americanus L3 larvae from fecal matter without much effort and in a clean manner.
The separation takes place naturally due to the positive phototaxis (movement towards light) which L3 larvae exhibit. So, while most of the process can take place in the dark, light is needed for at least a day before harvesting the larvae. This attracts them out of the fecal matter, towards the edge of the filter paper and into the drops of condensate which accumulate on the lid.
Petri dish. This can be made of glass or plastic. Glass petri dishes are re-useable and promote condensation more readily. There are several diameters available but they should not be too big. Around 60 mm is ideal.
Filter paper. Circular filter paper is available from laboratory equipment stores or eBay. A good and cheap alternative is unbleached coffee filter paper, which can be cut into circles using the lid of the petri dish as a guide to size. Two equally sized circles can be cut from entire coffee filters and placed on top of each other to match the thickness of a Whatman #3 filter paper.
Activated charcoal. This can be obtained from a pharmacy (expensive) or from the aquarium department of a pet shop (cheap). Any quality will do except the cheapest. In order to mix well with the fecal sample, the charcoal should be in powder form, and this can be produced by grinding the purchased charcoal in a blender. But be sure to keep the lid on the blender because a fine black dust is produced. Any powder residue left in the blender can easily be flushed away with water. Pre-powdered charcoal can be bought from a soap maker's supply store.
Pasteur pipettes and/or syringes (needle not required). These can be obtained from a pharmacy.
Test tubes. These are available from laboratory equipment stores, eBay or even scrapstores. I use 50 ml self- standing centrifuge tubes as an alternative. These are more difficult to find but they have a lid and stand on their own.
Heating mat. Any low wattage reptile heating mat is adequate.
Thermometer. Any thermometer capable of taking a measurement near the petri dish will suffice. I use a digital one with separate probe and min/max memory.
Cut the filter paper into a circle (the lid of the petri dish has roughly the right diameter) then wet the paper so that it is thoroughly soaked but not dripping. Then use the wet paper to line the petri dish so that it covers the entire bottom and side walls of the dish. The paper can touch the lid of the petri dish but it mustn’t prevent the lid from closing fully.
Mix the fecal sample with some powdered, activated charcoal, adding water if necessary. How much water will be required will depend on the consistency of the feces. For a sample with a normal consistency, a good starting point is a mixture with a feces/charcoal/water ratio off 1/1/1. Extra water or charcoal can then be added to create a mixture that spreads slightly but doesn’t run to fill the entire petri dish. Put this mixture onto the wet filter paper in the middle of the petri dish.
Close the lid of the petri dish and place it on the heating mat, then check the temperature because some mats get too warm. If this happens, you can place paper towels between the heating mat and the petri dish.
The ideal incubation temperature is said to be between 23°C (73°F) and 30°C (86°F), although one user of this method has found that a room temperature of 20-22°C (68-72°F), without a heat source, works well for him,  and another grower has had great success at 18°C (64-65°F).  At the lower end of the range, the larvae might be ready after 12-14 days and at the higher range after only 4-5 days. Larvae incubated at a lower temperature will have greater longevity.
No maintenance is required. Mold may form on the sample but this will not affect the culture.
The harvest can begin between 4 and 14 days, depending on incubation temperature. It’s better to wait a few extra days rather than go through the harvesting process only to find no larvae.
Expose the Petri dish to light for a day before collecting the larvae. This will encourage the larvae to climb. Any light source will do, but be careful with conventional lamps, especially those fitted with tungsten bulbs, since these give off heat and may overheat the culture.
Remove the lid, prop it at an angle on the dish and rinse the condensate off with a little water. To do this, fill a Pasteur pipette with either dechlorinated tap water or mineral water, or draw up 1-2 ml into a syringe. Squirt the water onto the raised end of the petri dish lid so the water takes the moisture drops down in the lid. Repeat this a few times to make sure everything is flushed off the lid. Suck up the rinse water using the pipette and put the collected water into a test tube or similar receptacle with a tapered bottom. I use self-standing centrifuge tubes. Now prop the petri dish at an angle on the lid and rinse the vertical and horizontal sides of the filter paper. Make sure the water only gets onto the filter paper and not between the dish and filter paper. Working systematically around the petri dish, rinse one section at a time, so the fecal sample in the middle remains untouched by the water. Then collect the water from the bottom of the petri dish, as well as any water left between folds of the paper at the top and the dish, and add this to the test tube.
Leave the collected water in the test tube for a day to allow the larvae to settle to the bottom or, in order to speed up the sedimentation of the larvae, a centrifuge can be used for 10 minutes at 1500 RPM. Then use a Pasteur pipette to collect a small sample from the bottom of the tube and check this under the microscope for L3 larvae. (Larvae that are not moving are not necessarily dead. They conserve energy until they find some skin. )
The culture can be re-moistened and the collection repeated daily until the there are no more larvae left to collect.
Any larvae that are not used immediately can be stored in the test tube for several weeks if they are kept within the ideal temperature range, in the dark - or at least out of direct sunlight - and provided with an adequate supply of oxygen.
One user has found that, if the 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 54ºF (12ºC) and 70ºF (21ºC) are unlikely to be a problem. 
The larvae do need oxygen, which can become depleted by higher temperatures and by bacterial activity in the storage tube. The latter can be resolved by opening the lid every few days and then gently shaking the tube. 
Even when kept in perfect storage conditions, the larvae will inevitably become progressively weaker as a result of the depletion of their fat stores.
This method delivers fairly clean larvae in comparison to the Harada-Mori culture technique (see download link) in which the water can get fairly dark and polluted with micro-organisms. The yield using the petri dish method depends on the size of the fecal sample and is comparable to that obtained using a Harada-Mori culture.
If required, the larvae can be cleaned further by using antibiotics and antifungals. The fecal sample can also be treated with antifungals to avoid mold growth. These topics will be considered in a future update.
- An Agar Plate Method for Culturing Hookworm Larvae: Analysis of Growth Kinetics and Infectivity Compared With Standard Coproculture Techniques
Users of a different incubation method have shared a number of valuable insights into hookworm culture that might be useful to those using this method by Steven.
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 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 detailed method by Alana. (This method provides information about every aspect of hookworm incubation.)