How to properly care for your stick and leaf insect eggs: A complete and comprehensive guide

One of the greatest joys of keeping exotic invertebrates is witnessing their fascinating life cycle. For me, there is nothing more rewarding than caring for eggs, watching them hatch, seeing the nymphs grow through each moult, and finally reaching adulthood to lay their own eggs. The cycle then proceeds to repeat.

The egg, otherwise known as ova, is a big part of this lifecycle due to the short-lived nature of invertebrates. Most species will spend nearly a third of their entire lifecycle developing within an ova.

But how should you go about keeping and incubating your stick and leaf insect eggs (ova) to achieve the best hatching success rate?

This is one of the most common questions among invertebrate keepers and finding a definitive answer online can be challenging. While there are many different techniques and methods for keeping, incubating, and hatching ova, some approaches prove more effective than others. Here, I will cover the common techniques used and the method which has worked most successfully for me across the vast majority of species I’ve kept.

Please note that this article provides standardised care information, as specific species will have different requirements. Some species are more tolerant while others are pickier about their conditions. Furthermore, some species naturally have low hatch rates – in these cases it may not be your care of the ova that is the issue. Finally, we need to be aware that phasmids have different egg-laying behaviours: some species bury their eggs in substrate, others glue them to food plants, and many simply drop them to the ground with an abdominal flick. This article focuses on the care of ova from the latter method which is the most common egg-laying behaviour.

The first aspect of stick and leaf insect ova care is choosing an appropriate incubation container. Key factors to consider regarding this incubation container are the size, ventilation, and ease of monitoring.

Size: The size of the incubation container should reflect and be appropriate for both the size of the hatching nymphs and the number of ova being incubated. A larger container is necessary when incubating either a high number of ova or ova from larger species. For example, the phasmid species Haaniella dehaanii produces extremely large eggs and consequently very large nymphs, necessitating a larger container. An inadequately sized container, whether due to nymph size or quantity, can lead to stress among newly hatched nymphs and may result in limb loss. While there's no strict rule for container dimensions, use your best judgment for your specific situation and species.

Ventilation: Proper ventilation is crucial for successful ova incubation. The incubation container must have small air holes that allow airflow while being small enough to prevent newly hatched nymphs from escaping. Adequate ventilation prevents stagnant conditions and, critically, helps prevent mould growth – one of the primary causes of embryo death and ova failure. Persistent mould growth often indicates insufficient ventilation and a sign that additional air holes should be added. Beyond mould growth ventilation also helps with gas exchange and the regulation of humidity and temperature. I recommend including bilateral ventilation in your incubation container, meaning air holes on either side. This allows for optimal airflow.

Ease of monitoring: Being able to easily monitor your ova is important for tracking development and identifying any potential issues such as mould growth. Therefore, I always recommend using a clear or transparent container as this allows you to observe the ova without disturbing them. This makes it easy to see both newly emerged nymphs and any signs of problems that may need addressing.

Based on these requirements, I strongly recommend using live food tubs – the type used for selling crickets, mealworms, and fruit fly cultures. These containers are ideal as they are cost-effective, (especially when reused or purchased in bulk), feature appropriately-sized ventilation holes (considering they are made for small invertebrates in mind), and are made of clear plastic for easy monitoring. Additionally, they come in various sizes to accommodate your species’ requirements. I personally use 220ml live food tubs. These measure ~10cm in width, 15cm in length and ~6cm in height.

That said, other containers can be used for ova incubation, including but not limited to takeaway containers, food tupperwares, and plastic or deli cups. However, these alternatives require additional preparation, such as adding ventilation.

Once you have a suitable incubation container, the next step is to choose the incubation “method” for your ova. This method primarily refers to your choice of substrate. The sections below aim to outline these methods.

The paper towel method is the simplest most common approach to incubating stick and leaf insect ova. Despite its simplicity, it can be highly effective when done correctly.

This method involves placing a single layer of paper towel, kitchen roll, or toilet paper at the bottom of your incubation container. While newspaper can be used, it’s not recommended as eggs are much harder to distinguish against its surface. Paper towel provides the best visibility for monitoring your ova.

To set up:

1. Cut the paper towel to fit precisely in the bottom of the container – avoid excess material that rides up the sides or requires folding, as this can create gaps where newly hatched nymphs can hide (and go unnoticed) or become trapped, resulting in unnecessary deaths.
2. Lightly spritz the paper towel with a mister until it is damp but not soaking wet. You can test this by squeezing the paper towel – if water drips out, it’s too wet.
3. Place the ova on the dampened surface, spacing them apart to prevent mould from spreading between the ova should it develop.
4. Re-spritz the paper towel whenever it dries out, taking care to remove the ova first and avoid spraying water directly onto them.

While this method is straightforward, it has two main drawbacks. First, paper towel is highly susceptible to mould growth because it's made of cellulose fibres – an organic material that mould can break down and use as food. Furthermore, unlike some natural substrates, paper towel has no antifungal properties or natural defences against mould. As a result, you'll need to replace the paper towel whenever mould appears, which can be as frequent as twice a week, making maintenance quite demanding. Second, paper towel dries out rapidly as it spreads moisture thinly rather than retaining it deeply. This means you'll need to spritz more frequently than with other substrates to maintain appropriate humidity levels, adding to the required maintenance.

The paper towel method is more than sufficient for successful hatch rates in common phasmid species such as the Indian stick insect (Carausius morosus) and the Macleay’s spectre stick insect (Extatasoma tiaratum).

The soil-based method is another common approach to incubating stick and leaf insect ova, using substrates like coco fibre, topsoil, or peat instead of paper towel.

This method involves placing a layer of substrate, ~2-3cm in depth, at the bottom of your incubation container.

To set up:

1. Place an even layer of soil (or alternative substrate) in the bottom of the container.
2. Lightly spritz the soil with a mister until it is damp but not soaking wet.
3. Place the ova on the dampened surface, spacing them apart to prevent mould from spreading between the ova should it develop.
4. Optionally, add a thin layer of sphagnum moss over the ova. This provides a surface for newly hatched nymphs to grip, helping them pull themselves free.
5. Re-spritz the soil whenever it dries out, taking care to remove the ova first and avoid spraying water directly onto them.

The soil-based method offers better moisture retention and requires less frequent maintenance than the paper towel method. However, I still do not recommend this method for several reasons. A major point is that soil typically contains organic materials that readily decompose, creating conditions that promote harmful bacterial and fungal growth which can damage developing embryos within the ova. Additionally, the dark substrate colour makes monitoring the ova and successful hatches difficult. Other challenges include increased difficulty to clean and replace the substrate when mould develops, harder to visually gauge moisture levels, and using soil can lead to uneven moisture distribution. Finally, there is also the risk of chemical residues if the substrate is sourced from non-specialised suppliers.

The suspended incubation method is a more specialised approach for incubating stick and leaf insect ova, using an elevated platform to suspend ova above a water source. This method provides humidity without directly contacting the ova.

To set up:

1. Fill a container or plastic cup with ~5cm of water.
2. Position a mesh or a ventilated tray about 1-2cm above the water level, ensuring the mesh holes are small enough to prevent the ova from falling through.
3. Place the ova on the surface of the mesh or tray.
4. Cover the setup with a second container that fits seamlessly over the first, leaving no gaps. This can be achieved by using two plastic cups – one larger, which holds the water, and one smaller, which fits securely on top. Together these form the incubation container. Alternatively, if the first container is tall enough, you can cut three sides of its top portion to create a hinged lid above the mesh or tray.
5. Ensure the top container or section has sufficient ventilation to prevent excessive humidity buildup.
6. Monitor water levels in the base container and top up as needed.

This method is very effective at preventing mould growth by providing good air circulation and avoiding direct contact with the substrate. Notably, this method has achieved positive results, demonstrating 100% trouble-free hatching for phasmid species like the giant lime green stick insect (Diapherodes gigantea) and the metallic stick insect (Achrioptera manga). However, this setup requires significant DIY to get it right and functioning. Furthermore, it can struggle to maintain adequate humidity for species needing consistently high levels, as water tends to evaporate from the base and pass straight out through the top ventilation.

With that said, for most species, if you have the time and materials, this can be an excellent incubation method.

The final method I will cover and the one I recommend is the vermiculite and plastic bottle cap method. This method is a highly effective, low-maintenance approach for incubating your stick and leaf insect ova.

The beauty and strength of this method lies in the use of vermiculite as the substrate. Vermiculite is a biologically inert material made from expanded mica, which naturally resists mould while also absorbing moisture helping to maintain high humidity levels. This offers superior mould prevention compared to paper towels or soil.

To set up:

1. Add a 3-5cm layer of vermiculite to the bottom of the container. Vermiculite comes in different particle sizes, from fine to coarse. Coarse vermiculite is generally preferred for ova incubation as it absorbs moisture without becoming overly compact, preserving airflow.
2. Lightly spritz the vermiculite with a mister until it is damp but not soaking wet. Despite its properties vermiculite is not completely exempt from developing mould. In rare cases over-wetting can encourage mould.
3. Place a clean, dry plastic bottle cap (such as from a milk carton, water bottle, or similar container) on top of the damp vermiculite layer. In cases where mould persists to develop, using a taller bottle cap with higher sides can help by creating a small barrier.
4. Arrange the ova within the bottle cap, optionally mixing in a small amount of dry vermiculite to further reduce mould risk and provide a gripping surface for newly hatched nymphs. This dry layer also helps absorb any condensation that may form on the cap.
5. When additional humidity is needed, simply remove the bottle cap, mist the vermiculite, and replace the cap with the ova.

I have personally achieved the highest hatching success rate across a wide range of phasmid and phyllium species using this method. Vermiculite’s moisture retention, mould resistance, and minimal maintenance requirements make it forgiving for new keepers whilst producing reliable, effective results for more experienced keepers.

So far, we have considered the incubation container and the incubation “method”. Now, let’s focus on the optimal conditions for incubating your ova, which will be divided into three key aspects: humidity, temperature, and lighting.

When determining the conditions for ova, it's essential to reflect on their natural habitat. I aim to mimic these conditions as closely as possible. As mentioned at the beginning of this article, the care guidelines are standardised for all phasmid and phyllium species, so these conditions should be adapted to the specific environmental needs of the ova you are caring for. Consistency is also critical; drastic fluctuations in temperature, humidity, or lighting cycles can be detrimental to the ova.

In my experience, the optimal humidity for most species’ ova falls within the 60-80% range. While we frequently encounter these percentages, what do they actually signify in terms of humidity? A humidity level of 60% means that the air contains 60% of the maximum moisture it can hold at a given temperature, essentially reflecting the air's capacity for moisture. However, this humidity does vary by species. For instance, Extatasoma tiaratum, a species native to Australia, thrives in much drier conditions and does not require nearly as much humidity for successful hatching. You can monitor humidity levels using a hygrometer.

To achieve and maintain a humidity of 60-80%, I spritz the vermiculite substrate with water approximately three times a week. This is especially critical when nymphs begin to hatch; excessively dry conditions can hinder their ability to emerge from the ova. In such cases, you may notice nymphs losing limbs or getting stuck in the ova capsule, which significantly reduces their chances of survival due to impaired mobility. If you miss a couple of humidity sprays, don’t worry – most ova can withstand dry periods of one to two weeks. However, prolonged dryness can lead to embryo death.

Additionally, it is important to use chlorine-free water for moistening the substrate, as chlorine can penetrate the ova and damage the embryo. To prepare chlorine-free water, let tap water sit for 24-48 hours to allow the chlorine to evaporate.

A consistent temperature is crucial for ova incubation, so it's important to minimise fluctuations; a variation of a few degrees is acceptable. I keep my ova at a consistent temperature of 23 °C (73.4 °F), but a temperature within the range of 20 °C (68 °F) to 26 °C (78.8 °F) is also ideal. to A mild nocturnal temperature drop is not a bad thing and can actually be beneficial as it mimics natural temperature cycles. For most species room temperature is generally sufficient, making additional heating unnecessary. I recommend avoiding heat mats near incubation containers, as they can lead to substrate desiccation and pose a risk of overheating the ova. Temperature levels can be easily monitored using a thermometer.

A natural photoperiod (day-and-night light cycle) is optimal for incubating and hatching ova, again replicating what the ova would experience in their natural habitat. I provide 12 hours of light and 12 hours of darkness, give or take a few hours on either side. The light cycle plays a significant role in determining the timing of hatching. Different species of nymphs tend to hatch at various times of the day, and they rely on this cycle for their emergence. In my experience, most nymphs hatch in the morning, typically between 9 AM and 12 PM. While research on hatching timing is limited, there is evidence that ova respond to light cues to decide when it is the right time to hatch. You can use either natural or artificial lighting, as both seem to have similar effects on the hatching process.

In concluding this article, it makes sense to address the biggest “enemy” of ova: mould. We have discussed trying to prevent mould extensively, but how much of a threat does it truly pose

Firstly, its important to recognise that mould is challenging to avoid, regardless of the incubation “method” employed. It is always best to catch mould early, as if left unchecked it can proliferate and become detrimental to your ova. For this reason, I recommend checking the ova daily, whenever possible.

If you notice mould has developed on the substrate, the solution is straightforward: simply remove or clean the substrate. However, if mould appears on the ova themselves, more caution is required. Gently remove the ova from the container and attempt to wipe off the mould with a clean paper towel or soft brush. If the mould growth is extensive, it's advisable to discard the affected egg to protect the health of the others.

Some sources state that springtails (Collembola) can be added to the incubation container to help control mould growth. These tiny, wingless arthropods feed on mould and can assist in managing its presence. However, this practice is somewhat controversial, as it is uncertain whether springtails might also harm or feed on the ova themselves. Personally, I tend to avoid using springtails due to this unknown risk.

It's worth noting that many people fear even the slightest presence of mould. In reality, a small layer of mould typically does not significantly impact the ova or embryo's health – do not panic! Harm occurs only when mould overwhelms the capsule and infiltrates it. Therefore, while mould should be monitored, it shouldn't be viewed as an overwhelming threat.

In this article, I have shared my insights on effectively caring for stick and leaf insect ova, highlighting various incubation techniques and optimal environmental conditions. As you embark on your own journey in keeping exotic invertebrate ova, remember that you will discover your own personal favourite methods, conditions, and maintenance routines that best suit your lifestyle and situation. There’s no one-size-fits-all approach to ova care and this article is intended as guidance to help you find what works best for you.

Thank you for taking the time to read.