Abstract
Several alternative approaches have been proposed to address the need for sustainable protein sources within our existing food and feed systems. Insects are regarded as a promising alternative, which has led to increased attention from researchers worldwide. Acheta domesticus, the house cricket, is considered a potential insect species for industrial production and various applications due to its high nutritious value. Since it is expected that A. domesticus and its applications will continue to rise as an important field of study, the standardisation of production techniques are needed. Nowadays, variation in the measurement of parameters and differences in experimental design limit the comparison among studies and, therefore, the ability to build upon existing knowledge. By identifying gaps in current protocols and providing suggestions on rearing practices and reporting, this paper aims to take the first step towards standardisation of the production and characterisation of Acheta domesticus for research purposes. This initiative primarily focusses on research practices evaluating the impact of feed and environmental conditions on the performance of house crickets.
1 Introduction
The growing concern regarding the sustainability of the current food and feed systems has led to the search for more sustainable protein sources. A promising approach that gained much interest lately is the use of insects in food, feed and technical applications. Insects are able to efficiently convert biomass to valuable components and have the potential to be produced more sustainably than current livestock (Bessa et al., 2017; Oonincx et al., 2010, 2015; van Huis, 2013; van Huis and Oonincx, 2017). Insects are a rich source of nutrients such as protein, fatty acids, vitamins and minerals (Magara et al., 2021; Rumpold and Schlüter, 2013; van Huis, 2013). Entomophagy, or the practice of consuming insects, is gaining traction as a viable solution to the challenges facing traditional livestock farming, such as the high food demand, and various species are already consumed by humans in many parts of the world (Jongema, 2017; van Huis, 2013).
One of the insect species considered promising for food applications is the house cricket, Acheta domesticus (Jensen et al., 2017). In the European Union (EU), insects are considered a Novel Food and therefore are subjected to the Novel Food legislation, i.e. Regulation (EU) No 2015/2283. In 2022, a novel food dossier was approved for the house cricket (2022) (frozen, dried and ground) (Fernandez-Cassi et al., 2018; IPIFF, 2021). Crickets are a highly nutritious source of food, with a protein content ranging from 56% to 67% on dry matter (DM) basis (Bawa et al., 2020; Collavo et al., 2005; Harsányi et al., 2020; Oonincx et al., 2015). One of the benefits of house crickets is their lower environmental impact compared to vertebrate livestock (Kuo and Fisher, 2022; Marcucci, 2020). For example, crickets emit 0.05 g carbon dioxide per kg body mass per day, whilst pigs produce 30.46 g per kg per day and cattle 16.52 g per kg per day. Regarding ammonia emissions, crickets produce 5.4 mg ammonia gas per kg body mass per day, while the pigs produce, respectively, 91.2 mg/kg per day and cattle 53.4 mg/kg per day (Oonincx et al., 2010; Philippe and Nicks, 2015; Rzeźnik and Mielcarek, 2016).
The potential of the house cricket as an alternative food source has led to increased attention from entrepreneurs and researchers worldwide. The number of publications generated on ‘Acheta domesticus’ has increased over the last decade, with 31,6% of all studies since 1960 published in the last 5 years alone (Figure 1).
Number of publications published during the last 10 years using ‘Acheta domesticus’ as keyword.
Citation: Journal of Insects as Food and Feed 10, 7 (2024) ; 10.1163/23524588-00001042
Today, the growing interest by researchers for the mass production of A. domesticus has led to knowledge gain on the insects’ behaviour and requirements. For instance, it currently is well known that many factors influence A. domesticus performance, such as climate conditions in which the insects are reared (Busvine, 1955; Clifford et al., 1977), their diet (Bawa et al., 2020; Kuo and Fisher, 2022; Oloo et al., 2019; Oonincx et al., 2015; Orinda et al., 2017; Sorjonen et al., 2019), their housing, and density (Clifford et al., 1977; Morales-Ramos et al., 2020; Tennis et al., 1977). However, as the application of insects in food and feed is a relatively new concept in the western society, many research questions still need to be addressed in order to optimise the production and application of house crickets (Dobermann et al., 2017; Jensen et al., 2017; Kuo and Fisher, 2022; Lundy and Parrella, 2015). Consequently, cricket production and processing will most likely further become an important field of study.
Considered as one of the most important gaps in insect research is the lack of standardised methods among studies. For instance, the differences in measurement of rearing-relevant parameters hamper comparison across publications. Studies may refer to ‘growth’ as ‘yield’, ‘weight gain’, ‘final or end weight’, ‘dry body mass’, ‘maximal weight’, etc. The same accounts for crickets’ feeding efficiency, which may be referred to as ‘feed conversion ratio (FCR)’, ‘bioconversion efficiency’, ‘efficiency of conversion of ingested food (ECI)’, ‘ingested food’, etc. Often, raw data is not presented, making it impossible to recalculate to common parameters. Furthermore, various studies may implement different life stages at the start of the experiment as well as different harvesting strategies and times (Bosch et al., 2020; Kuo and Fisher, 2022; ValuSect, 2022; Van Peer et al., 2021).
Regarding chemical analyses, different methods can be used to determine the insects’ composition. Different analysis techniques lead to differences in data measurements and often the methods used are not clearly described. The same accounts for the pretreatment of samples prior to analysis, which also impacts the results (Inácio et al., 2021).
It is clear that there is a need for guidelines and methods on the rearing and characterisation of A. domesticus for research purposes in order to provide the sector with relevant data and knowledge to build upon. In 2020, Bosch et al. (2020) emphasised the need for guidelines on black soldier fly research and described the first basis for a protocol. To date, this has not yet been the case for A. domesticus. Several rearing methods have been described previously, e.g. Clifford et al., 1977; Clifford and Woodring, 1990; Patton, 1978, however, many new insights have been obtained from the intensified research in recent years.
This paper (1) summarises the literature on optimal rearing methods for A. domesticus in research settings, (2) aims to identify gaps in current protocols and (3) provides specific suggestions for methods and reporting that were validated by the authors in various research projects prior to the writing of this paper (e.g. Van Peer et al., 2023). This paper represents an initial step towards standardisation of house cricket rearing and reporting for research purposes, more specifically for the performance of house cricket feed experiments and the optimisation of environmental conditions.
2 Environmental conditions
Insects are ectothermic, hence requiring certain climate conditions for satisfying growth and development (Bjørge et al., 2018). Unless it is part of the research question, it is therefore highly recommended to apply climate conditions defined as optimal for cricket rearing (Clifford et al., 1977; Patton, 1978). Several authors report an optimal temperature for A. domesticus between 30 °C and 35 °C (Clifford and Woodring, 1990; Ghouri and Mcfarlane, 1958; Roe et al., 1980). Temperatures below 25 °C have shown a negative impact on crickets’ growth and development (Booth and Kiddell, 2007; Busvine, 1955; Roe et al., 1980). For instance, crickets reared at 28 °C reached maturity 70 days earlier than crickets reared at 25 °C (Booth and Kiddell, 2007) and lowering the rearing temperature from 30 °C to 23 °C resulted in development times 3 times higher for male and 4 times higher for female crickets (Busvine, 1955). Crickets grown at a temperature of 21 °C were found unable to thrive (Booth and Kiddell, 2007). On the other hand, breeding temperatures slightly above 35 °C (38 °C) seem to cause high mortality (Ghouri and Mcfarlane, 1958).
It has been shown that even small differences in climate conditions, can lead to significant differences in growth parameters. For instance, increasing the temperature from 30.5 °C to 31 °C, resulted in a 10-day shortening of the life cycle. Consequently, the optimal temperature for cricket rearing based on the shortest development time was defined as 35 °C (Clifford et al., 1977). However, Clifford and Woodring (1990) recommended a rearing temperature for A. domesticus of 30 °C. This suggestion was based on a compromise between human comfort, feasibility and house cricket performance. For instance referring to the danger of high mortality accruing at small temperature fluctuations above 35 °C. In addition, dry weight gain showed no statistically significant differences at rearing temperatures of 30 °C and 35 °C (Clifford and Woodring, 1990; Roe et al., 1980).
The relative humidity in which house crickets are kept is also an important factor, especially to guarantee the survival of eggs and young crickets (Ghouri and Mcfarlane, 1958). Reported by Clifford (Clifford and Woodring, 1990), older crickets require a lower relative humidity (i.e. 50%), as higher humidities might kill crickets from the 4th through last instar nymphs and adults. To incubate eggs prior to experiments, it is recommended to maintain higher relative humidity levels while limiting the risk of infestation from mites and mould. Setting the relative humidity at 60% in the incubation chamber is suggested (Clifford and Woodring, 1990; Ghouri and Mcfarlane, 1958).
Crickets are nocturnal insects, preferring to lay eggs in darkness. They avoid open, lighted spaces and seek shade for resting (Cymborowski, 1973). It has been reported that applying a photoperiod of 12:12 or 14:10 hours light-dark is important to support their circadian rhythm (Clifford and Woodring, 1990). This is based on induced activities of crickets when a photoperiod is present (e.g. locomotory activity) (Bertram and Bellani, 2002; Cymborowski, 1973; Nowosielski and Patton, 1963). In natural conditions, the house cricket is mostly active between 19.00-24.00 h. In natural light conditions, it shows a distinct rhythm, with highest activity occurring immediately after dusk (Cymborowski, 1969). During daytime A. domesticus is mostly hidden in its retreat (Kieruzel, 1976). This might suggest the feasibility of growing crickets in the absence of light, especially considering the successful completion of full breeding cycles at our facilities where A. domesticus was reared in complete darkness.
In conclusion, it is suggested to set the climate conditions at 30 °C and 50% RH and rear crickets in darkness during the experiment and for the parental population.
Considering the influence of environmental factors on the growth and performance of insects, these climate conditions, including temperature, humidity and photo regime, should always be reported. Moreover, it is recommended to perform experiments with crickets in a climate-controlled room, incubator or cage, which provides automated control and minimises fluctuations. Additionally, tracking climate conditions using loggers is important to intervene in case of fluctuations or to exclude climate as a variable.
3 House crickets’ (control) diet
Another factor affecting the growth and development of house crickets is their diet (Bawa et al., 2020; Kuo and Fisher, 2022; Oonincx et al., 2015; Orinda et al., 2017; Sorjonen et al., 2019). Crickets’ development time, survival rate, growth rate and feed conversion efficiency depend on the nutritional composition of their diet. For instance, high (30.5% on a DM) and medium (22.5% DM) crude protein diets perform significantly better in terms of cricket yield, growth and development compared to low protein diets (15% DM) (Sorjonen et al., 2019). Diets containing low protein contents lengthen the crickets’ development time and result in higher mortalities compared to higher protein diets (Oloo et al., 2019; Oonincx et al., 2015; Sorjonen et al., 2019). Several studies have reported that a protein content of 20-30% supports the growth of crickets (Clifford and Woodring, 1990; Oloo et al., 2019; Orinda et al., 2017; Patton, 1967; Woodring et al., 1979).
In Patton’s study (1967), four diets were identified to support the growth and survival of house crickets. The carbohydrate content of these diets ranged from 32-47% on a DM basis, while the fat content varied between 3.2 and 5.2% (Patton, 1967). Also in other studies higher levels of carbohydrates have been found to improve A. domesticus’ survival rates, weight gain, and to accelerate development (Orinda et al., 2017; Vaga et al., 2020a,b).
It has been shown that crickets successfully grow on commercial animal feeds such as chicken, dog- and rabbit feed (Nakagaki and Defoliart, 1991; Patton, 1963). Chicken feed is a standard feed widely used for growing crickets (Akinyi Orinda et al., 2017; Clifford et al., 1977; Halloran et al., 2017; Hanboonsong and Durst, 2020; Orinda et al., 2017; Quek et al., 2019). However, it has been suggested to grind the feed as crickets tend to be selective regarding the particle size, rejecting large particles which may contain essential materials (Clifford et al., 1977; Clifford and Woodring, 1990; Patton, 1963; Tennis et al., 1979). Furthermore, increased access to water results in higher cricket growth rates (McCluney and Date, 2008).
Crickets can be provided with different moisture sources. For instance, commercial chicken watering systems modified with sponges (Clifford and Woodring, 1990; Cortes Ortiz et al., 2016). However, this system might be less preferred due to hygiene concerns as crickets are prone to diseases (e.g. Densovirus) that can cause significant mortality within the population (Eilenberg et al., 2015; Lecocq et al., 2018). Solid sources of moisture such as fruit and vegetables and agar cubes can also be used (Cortes Ortiz et al., 2016; Coudron et al., 2022a,b; Deruytter et al., 2021). Agar cubes have the advantage that the nutritional composition is well known and stable, containing mostly water and only few other elements that might influence experimental outcomes. As agar is a solid moisture source, it excludes the risk of crickets drowning.
Set-up for cricket rearing using egg cartons as hiding spots.
Citation: Journal of Insects as Food and Feed 10, 7 (2024) ; 10.1163/23524588-00001042
In the context of the house crickets’ (control) diet, we suggest using ground chick feed (20-30% crude protein) and agar as moisture source (both ad libitum) as this has demonstrated satisfactory results in terms of cricket growth and performance. Regarding reporting, it is important to mention the supplier of the feed but more importantly the nutritional composition of the feeds used, as the feed properties may vary by supplier and even batch (Broeckx et al., 2021; Van Peer et al., 2021). Reporting the dietary nutritional value should include at least the proximate composition, namely dry matter, crude protein, crude fat, ash content and crude fibre.
4 Housing
Different housing options are available for rearing crickets in laboratory settings such as terrariums, crates (e.g. 40 × 60 cm) and cages of various sizes (Collavo et al., 2005; Oonincx et al., 2015; Patton, 1978). Therefore, it is important that the type of housing, including material and dimensions, are reported. Regarding the material of the rearing crate, crickets cannot climb smooth surfaces (e.g. plastic, metal) (Clifford and Woodring, 1990; Mukherjee and Mukherjee, 2022). However, note that glass containers can cause humidity problems due to condensation and might therefore be less suitable (Clifford et al., 1977). These types of housing should be placed in an incubator or climate controlled room to minimise the effect of climate fluctuations. Automated systems are also a possibility. These are usually equipped with temperature control, light and ventilation within the system, so that the entire rearing chamber does not have to be monitored (in contrast to housing in an incubation chamber). In case of using automated cages, the technical data should be reported in publications such as ventilation speed, air changes/hour, volume of the rearing chamber/cage and set climate conditions.
As crickets avoid open spaces, hiding places should be present in their housing (Clifford and Woodring, 1990; Kieruzel, 1976). Egg cartons can be used as hiding spots and as a way to increase usable volume (Figure 2) (Cortes Ortiz et al., 2016; Mukherjee and Mukherjee, 2022).
Factors such as density also affect crickets’ growth. For instance, Morales-Ramos et al. (2020) found differences in A. domesticus performance when feeding identical diets to small and large groups. Tennis et al. (1977) investigated the effects of 3 different densities on crickets’ growth and survival and found that cricket’s mortality increased and individual weight decreased with increasing density. According to Patton (1978) a minimum space of 2.5 cm2/cricket leads to the best growth and survival. Higher densities lead to higher cricket mortalities. Therefore, it is suggested to apply a density of 0.42 nymphs/cm2 of the tray ground surface. This is also supported by a recent study (Mahavidanage et al., 2023). Laboratory-scale experiments involving only a single cricket may be pertinent to certain research objectives. However, when performing practice-oriented research, it is advised to conduct experiments with the suggested density (multiple crickets) to enhance the translatability of the results to the sector.
5 Origin of the crickets and parental population
It has been shown that differences in insect genetic background might lead to variation of results. For instance, it has been shown that different strains of Tenebrio molitor, yellow mealworm, resulted in variation in terms of larval performance. In the study of Rumbos et al. (2021), seven strains from different geographic origins were evaluated on larval growth and development when reared under identical conditions. It was concluded that larval end weight and development time as well as feed efficiency parameters were significantly influenced by strain. Regarding insect composition, differences in dry matter content as well as energy content were found. Similarly, Morales-Ramos et al. (2019) reported differences in terms of growth, survival and ECI between two strains of T. molitor; one strain being the result of 8 years continuous, artificial selection for larger pupal size, and one ancestral strain. The same was shown for the black soldier fly (Hermetia illucens). In the study of Zhou et al. (2013) it was demonstrated that the development and waste conversion of black soldier fly larvae differs among strains. A more recent study also reported clear strain effects (larval performance and composition) when feeding three different diets to 4 different populations of black soldier fly larvae (Sandrock et al., 2022).
Considering the above, it is important that in any study information on the origin of the insects is provided. Insects that have been reared for multiple generations in the laboratory, might respond differently to experimental feeds or conditions than insects collected in the wild; and insects artificially bred in a certain way, might develop different properties than insects reared differently. This is a consequence of inevitable (and not always intended) selection for certain parameters or adjustment to a certain environment or diet (Morales-Ramos et al., 2020; Wang et al., 2017).
If an experiment is conducted with a wild strain, this, as well as the location of collection, should be reported. If the wild strain is kept in the rearing facility for a few generations, also this should be mentioned as well as the number of generations. In case the insects were purchased from an insect breeder, it is suggested to report the name of the company and their location (Bosch et al., 2020). For insects that have been reared continuously in the research facility, the rearing of the parental population should be reported. This includes the climate conditions in which they are kept as well as the parental age and collection method and incubation conditions of the eggs. Specific suggestions regarding the latter are provided below.
Ideally, the parental population consists of adult crickets that are at least 10 days old, as studies have demonstrated that oviposition starts from this point onward (Clifford and Woodring, 1990; Murtaugh and Denlinger, 1985). As soon as the males start chirping, the colony is maturing and will start mating.
Specifications on the oviposition tray, such as the used substrate, should be reported. The oviposition tray (preferably 15 cm in width/diameter or more and at least 3 cm deep) is filled with a substrate such as peat (without fertilisers), which is moistened before use (30-35% DM). Female crickets will not oviposit their eggs unless a suitable, moist substrate is present that ensures survival of the offspring (Clifford and Woodring, 1990). It is advised to cover the oviposition substrate with a mesh (2 mm) in order to prevent crickets kicking the substrate (and eggs) from the tray (Figure 3). The oviposition tray should be changed at least every 72 hours to prevent dehydration of the substrate. At 30 °C, house crickets’ eggs require on average 13 days to hatch (Clifford and Woodring, 1990).
Oviposition tray for cricket rearing.
Citation: Journal of Insects as Food and Feed 10, 7 (2024) ; 10.1163/23524588-00001042
6 Insect performance parameters
Insect development time, survival, growth rate, and efficiency of converting food into biomass and chemical composition are commonly used parameters to evaluate insect performance (Bosch et al., 2020; Kuo and Fisher, 2022). The development time refers to the time necessary for the insects to grow to a specific point, such as when all feed is digested or when a certain percentage of adults are present, i.e. the duration of the experiment, if not a fixed time is used to end the experiment (Bosch et al., 2020). Evaluating the development time is a valuable method for assessing treatments, as non-optimal conditions often result in longer development times.
Survival of the crickets is estimated by determining the density at the start and at the end of the experiment. Determining the survival allows to detect potential harmful components or conditions present in the treatment. The survival is expressed in percentage.
It is recommended to determine at least these above-mentioned parameters to track insect performance and to determine treatment effects. Formulas of all measurements used to determine insect growth, performance and conversion efficiency should be reported.
As various methods are applied among studies to assess the conversion efficiency of feed by insects, this topic will be discussed in more detail below. Individual growth of insects will also be further addressed as, in our experience, the determination of this parameter is prone to errors.
Efficiency parameters
Efficiency parameters are particularly of importance during the performance of feed experiments. For other types of experiments, these parameters might hold less relevance. There are different formulas applied to determine the efficiency by which insects transform feed into biomass (Bosch et al., 2020; Kuo and Fisher, 2022). Determining these measurements allows to investigate the response of insects to different diets in terms of growth and performance (Scriber and Slansky, 1981). Efficiency parameters include feed conversion ratio (FCR), bioconversion efficiency (BE), efficiency of conversion of ingested food (ECI), efficiency of conversion of digested food (ECD) and waste reduction index (WRI).
Formulas to calculate the efficiency parameters described below are presented in Table 1.
Feed conversion ratio
The FCR can be defined as the amount of feed needed per unit of larval biomass increase (Bosch et al., 2020). The FCR can be determined using the formulas described by Scriber and Slansky (1981). According to the formula, the lower the FCR, the better the conversion.
FCR values are often used to compare feed efficiency between conventional livestock and insects, however, this comparison is questionable. For instance, factors influencing FCR encompass dressing percentage and carcass refuse and should be included to obtain comparable values. In the case of insects, the edible fraction is typically significantly greater than that of vertebrate livestock, with most of the insect being edible (up to 100%) (Marcucci, 2020; Nakagaki and Defoliart, 1991).
The FCR can be calculated on a dry, wet, and dry/wet base. FCR values based on wet matter might be challenging to compare, considering the large variations in the dry matter content of different diets and the produced insects. Therefore it is suggested to generally present efficiency parameters on DM basis (Bosch et al., 2020). However, the water intake of insects and vertebrate livestock may not always be included in the calculations (Marks, 1981; O’Meara et al., 2020). Comparing FCR values among insects and between insects and livestock should therefore be addressed with due caution.
Bioconversion efficiency
The bioconversion efficiency of the insects can be determined by using the inversed formula of FCR and is often used to compare feed efficiency among insects in experimental settings. As extension to the bioconversion efficiency, there is a formula to correct on the residual frass, a mixture of leftover food, faeces and insect residues which is not processed into biomass. This is called the BER, bioconversion efficiency corrected for residue. BE and BER are calculated in g DM (Bosch et al., 2020).
Efficiency of ingested/digested food
Some studies also use the ECI (efficiency of conversion of ingested food) as efficiency parameter, which is a variant of the bioconversion efficiency. The amount of ingested feed by the insects is estimated by the amount of feed given during the experiment and the amount of feed left at the end of the experiment (in g DM). Slow growth is associated with low ECI values (Scriber and Slansky, 1981; Waldbauer, 1968).
Another extension to this formula has been described, namely the efficiency of conversion of digested food (ECD). The ECD aims to correct only for the amount of faeces left at the end of the experiment, meaning that these should be separated from insect leftovers and unprocessed feed.
Important to note is that determining the ECI and ECD as efficiency parameters are not suggested for insects thriving in their substrate such as mealworms and black soldier fly larvae as separating the frass into fractions and determining the amount of leftover feed and faeces might lead to large errors (Bosch et al., 2020). As crickets don’t necessarily thrive in their substrates and feed systems are often used, it might be easier to determine these parameters for this species. However, keep in mind that this is only an estimate as crickets tend to move their feed throughout their crate.
Waste reduction
If relevant to the research (e.g. feed experiments with byproducts), the waste reduction index can be calculated using the formulas of Diener et al. (2009) and Rehman et al. (2017). The waste reduction index refers to the insects ability to reduce organic biomass.
Overview and formulas of efficiency parameters
Citation: Journal of Insects as Food and Feed 10, 7 (2024) ; 10.1163/23524588-00001042
Individual growth
It is advised to track the growth of the nymphs during the experiment. By doing this, and combined with initial and harvest weight, a growth curve can be constructed and the growth rate can be determined. For assessing growth, we propose using the subsampling method as described below.
Subsampling is used to estimate the average weight of randomised individuals. Assessing crickets’ growth on at least a weekly basis is advised to gather a sufficient amount of reliable data. Subsampling is done by collecting randomised individuals from the rearing crate. Due to the inherent size variability of the nymphs, a small subsample may result in large errors on the average weight estimate. Therefore, a subsample size of at least 100 individuals should be used. After collecting, the subsample is weighed exactly and every nymph present in the subsample is counted. The average weight of 1 nymph can then be calculated. As the nymphs are quite mobile, counting is preferably done by taking a proper picture of the collected nymphs. An example of such picture is shown in Figure 4. To limit errors on subsample data, it is recommended to take at least 3 subsamples per crate/cage and make calculations using the mean weight. The cricket counting process can be facilitated through the utilisation of artificial intelligence, wherein advanced algorithms can efficiently detect and quantify the crickets present in a given box based on images or videos. This innovative approach not only expedites the counting procedure but also enhances accuracy, providing a swift and precise assessment of the cricket population within the designated environment.
Example picture for cricket subsampling.
Citation: Journal of Insects as Food and Feed 10, 7 (2024) ; 10.1163/23524588-00001042
7 Start of the experiment
Given the vulnerability of young nymphs to handling (Hanboonsong and Durst, 2020; Vaga et al., 2020b), we propose initiating the experiment two weeks after hatching of the eggs, unless the focus of the experiment pertains to pinheads (e.g. hatching rate). This initial two-week period, is referred to as the ‘nursery period’. To ensure minimal age variability among the cricket population, it is advisable to collect hatchlings that are no more than 24 hours old.
Two week old crickets can then be collected for the experiment. To maintain uniformity within the cricket population, we recommend collecting all crickets intended for the trial in a single crate. Next, these crickets should be divided into different treatment groups with equal densities using the subsampling method described earlier. It is crucial to perform at least three subsamples to reduce errors and determine the coefficient of varience. If the coefficient of variance of the amount of crickets (per gram of sample) among the subsamples exceeds 10%, additional subsamples should be taken until the it falls below this threshold.
After the nymphs are collected and divided into treatments, they are incubated as previously described.
8 Harvest of crickets
At optimal growth conditions, it takes approximately 6 weeks from hatching of the eggs until nymphs reach maturity (Clifford and Woodring, 1990; Patton, 1978). Different harvesting times have been applied among studies. For instance, using a fixed time (Harsányi et al., 2020; Lundy and Parrella, 2015) as well as at a specific life or development stage (Magara et al., 2019) and at appearance of first adults (Oonincx et al., 2015; Vaga et al., 2020b). Using a fixed time to end the crickets experiment excludes certain variables and allows data among studies to be compared. For instance, development on a less suitable diet might be lower and ending the experiment when a certain percentage of the population has reached maturity requires an experienced eye. The latter was found to be subjective and high standard deviations have been detected (Van Peer et al., 2023). Fixed times to end the experiment are easier as they can be scheduled and are therefore recommended.
In addition, different methods can be used to harvest crickets. As an example, crickets can be manually harvested by utilising the egg cartons (hiding spots). Fit crickets tend to stay attached to these cartons, and can therefore easily be transferred (shaking off). Crickets can also be harvested by sieving, to separate them from their frass (Hanboonsong and Durst, 2020; Mukherjee and Mukherjee, 2022; Spranghers et al., 2021). As some harvest methods might lead to losses or the inclusion of deceased individuals, the method of harvesting should be reported.
9 Statistical analysis of growth and performance data
To be able to conduct proper statistical analysis, it is generally advised to apply at least 4 replicates per treatment when performing experiments. As described by Bosch et al. (2020), these replicates are typically carried out concurrently, utilising the same generation or population of insects. While this approach enhances the precision of treatment effect determination, it may inadvertently limit the generalisability of the data for insect breeders worldwide. Hence, it is advisable for future studies to incorporate replicates spanning multiple insect generations or populations.
Various methods can be employed to detect statistical differences among growth parameters. When dealing with normally distributed values, distinctions can be detected through an ANOVA test. The Tukey HSD test is a widely-used data analysis method that facilitates the comparison of means across multiple groups, enabling the detection of intergroup differences. If the variances differ from each other, Welch’s test can be performed, however, this test does not allow comparing the groups. Furthermore, the Tukey HSD test includes a correction for the number of group comparisons, reducing the likelihood of family-wise error rates or type I errors. The Kruskal-Wallis test, a non-parametric statistical test, is deployed to compare the medians of three or more independent groups. It serves as a viable alternative to the one-way ANOVA test in cases where normality hypothesis is violated or when sample sizes are too small to verify normality.
Due to the broad range of methods available, the method applied to analyse data should always be reported.
In the context of growth curves, it is essential to seek a suitable model that relates individual weight to time. Various logistic models have been proposed to assess the effect of insect growth parameters. Nijhout and Wheeler explored the potential application of the Gompertz growth model for holometabolous insects (Nijhout and Wheeler, 1996). Logistic models have also been applied to describe the growth of black soldier fly larvae in previous studies (Bekker et al., 2021; Sripontan et al., 2020). Mixed effect modelling might be an alternative, as this takes the longitudinal nature of growth data into account. This technique has been used by Coudron et al. (2022b) and Deruytter et al. (2021). Unfortunately, the formula used for models is not consistently reported in the conducted studies, making it challenging to compare data due to the wide array of models available. When addressing cricket growth, it is suggested to employ a biological growth model as opposed to a black box model. Utilising a biological growth model offers several advantages, such as providing a more reliable representation of real-world growth patterns, enhancing transparency and interpretability, and yielding more precise predictions of growth across diverse environmental conditions (Cavazzana et al., 2007; Fung et al., 2021; Loyola-Gonzalez, 2019).
10 Chemical analysis of house crickets
Determining the chemical composition of insects might be relevant for certain research questions. For instance, when performing feed experiments, it is recommended to at least determine the proximate composition of the crickets produced on these diets.
In order to be able to compare published study outcomes for analytical data, attention should be paid to the methods and sample preparations that were used. It is advisable to work with standardised methods and their sample preparation method. The used analytical methods should be reported or referred to fully understand the results. The International Organisation for Standardisation (ISO) collects standardised analytical protocols for different matrices. Standardised protocols for insects can be found as ‘foodstuff’.
The preparation method of samples significantly influences the analysis results. For instance, when studying the effect of the feed substrate on insect composition, crickets should be starved for a minimum of 24 hours after harvest prior to performing insect analysis. This fasting period ensures that their gut load, which represents their substrate, is emptied (Barker et al., 1998; Finke, 2002; Finke and Oonincx, 2013). This should provide the assurance that the nutritional components have been digested by the insect and are not retained in the gut.
To determine the proximate analysis, it is recommended to take at least 50 g of fresh sample. The preparation method depends on the specific analyses to be conducted. In the case of nutritional analyses, it is suggested to dry the samples at a maximum temperature of 60 °C or freeze-dry them, according to ISO 6498:2012 ‘Animal feeding stuffs: Guidelines for sample preparation’. This to preserve the integrity of the insect composition and temperature-sensitive components, such as e.g. amino acids or B-vitamins (Oonincx and Finke, 2021).
The termination method used on the insects, like blanching or freezing, should also be reported. This helps comparing study results of delicate components. Subsequently, samples must be homogenised in order to perform a reliable and correct analysis. Generally, samples are ground either manually or automatically and sieved through a 1 mm mesh according to ISO6498:2012.
Proximate analysis should include at least crude content of protein, fat (or ether extract), dry matter content, ash and crude fibre. Mineral composition and other compounds (e.g. vitamins, metals, etc.) or more detailed analyses (e.g. amino acid profiles and fatty acid composition) may also be relevant for some research questions. It is important to report which analytical methods were used and how results were calculated (Finke and Oonincx, 2013). For example, for determination of the crude protein content two different methods are commonly used. The Dumas and Kjeldahl methods are important in the quantitative determination of nitrogen content in chemical substances. The crude protein content is calculated based on the nitrogen content. Whereas the Kjeldahl method is the most frequently applied method over the years, nowadays the Dumas method is gaining increasing popularity in food laboratories thanks to its speed and reliability. Results obtained with the Kjeldahl method are usually a bit lower than with the Dumas nitrogen determination. The Dumas method measures all organic and inorganic nitrogen through combustion. The Kjeldahl method is not able to determine certain compounds like nitrates, nitrites and heterocyclic compounds. Both techniques are standardised and differences in total nitrogen content of insects are small (Thompson et al., 2002). Large differences are found based on the nitrogen (N) to protein (P) factor applied to calculate the crude protein content. As these techniques have been validated for protein determination for meat, eggs or milk products, the general N-factor 6.25 is overestimated for insects (Van Huis et al., 2013). The exoskeleton of insects is made of nitrogen-containing chitin fibres. Therefore, previous studies have determined more accurate N to P factors that can be used for insects. Factors that can be used for edible insects are 5.33 (Boulos et al., 2020) and 4.76 (Janssen et al., 2017), or more specific for house crickets 5.09 (Ritvanen et al., 2020). Nevertheless, accurately determining the crude protein content through this approach remains challenging. A more precise method involves analysing amino acids to calculate the true protein content. The amount of chitin in insect cuticle is very variable. Hard cuticles have a chitin content up to 70-85% while soft cuticles contain 15-30% of chitin. As a consequence, the chitin content between nymphs and adult crickets differs (Chapman, 1998). To assess the chitin content several analytical methods can be used, depending on the research topic (nutritionally or chemically). Nutritionally, chitin is a fibrous compound and methods for fibre determination can be used based on the level of in-depth study of its digestibility like: crude fibre content (CF), acid-detergent fibre (ADF), neutral detergent fibre (NDF) or dietary fibre content. When studying chitin content in insects with a more chemical approach, for example to transform it into chitosan for technical applications, more specialised methods like UPLC/FLR are used (Han and Heinonen, 2021).
Overview of suggested rearing factors and minimal data to be reported
Citation: Journal of Insects as Food and Feed 10, 7 (2024) ; 10.1163/23524588-00001042
(Continued)
Citation: Journal of Insects as Food and Feed 10, 7 (2024) ; 10.1163/23524588-00001042
11 Summary
Since insects have emerged as a sustainable alternative protein source, there has been a notable increase in scientific publications on their implementation in research. Insects have become a significant area of study nowadays, however, publications often lack comprehensive information, hindering effective comparisons. Often, the methods are inadequately described, and measurements for rearing practices and chemical analysis are not provided. In 2020, Bosch et al. (2020) described the first basis for a protocol on black soldier fly research, however,
comprehensive guidelines and procedures for conducting research on A. domesticus are currently still lacking (Kuo and Fisher, 2022). In this regard, this paper aims to provide a basis in the standardisation of house cricket research by identifying gaps in current protocols and providing specific suggestions for methods and reporting on cricket production and analysis.
Below (Table 2), we present an overview of the minimal data to be reported and the suggested rearing protocol for A. domesticus in research contexts, with a primary focus on experiments evaluating the effect of feed and environmental conditions on cricket performance. This overview can serve as a guidance during the performance of experiments involving A. domesticus and the subsequent reporting of methods and results.
Corresponding author; e-mail: meggie.vanpeer@thomasmore.be
Authors’ contribution
All authors provided input on sections within their expertise. M.V.P. drafted the work and undertook literature review on all sections with the help of S.B. and C.C. I.N. provided input on the section of chemical analysis. S.V.M. supervised and reviewed the work together with G.R.V. All authors have read and agreed to the published version of the manuscript.
Conflict of interest
The authors declare no conflict of interest.
Funding statement
This review was funded by Interreg NWE, grant number NWE1004, ValuSect-Valuable inSects; and co-funded by Vlaams Agentschap Innoveren & Ondernemen.
References
Akinyi Orinda, M., Oyoo Mosi, R., Awuor Ayieko, M., Anangwe Amimo, F. and Oginga Odinga, J., 2017. Effects of housing on growth performance of common house cricket (Acheta domesticus) and field cricket (Gryllus bimaculatus). Journal of Entomology and Zoology Studies 5: 1138-1142.
Barker, D., Fitzpatrick, M.P. and Dierenfeld, E.S., 1998. Nutrient composition of selected whole invertebrates. Zoo Biology 17: 123-134.
Bawa, M., Songsermpong, S., Kaewtapee, C. and Chanput, W., 2020. Effect of diet on the growth performance, feed conversion, and nutrient content of the house cricket. Journal of Insect Science 20: 10. https://doi.org/10.1093/jisesa/ieaa014
Bekker, N.S., Heidelbach, S., Vestergaard, S.Z., Nielsen, M.E., Riisgaard-Jensen, M., Zeuner, E.J., Bahrndorff, S. and Eriksen, N.T., 2021. Impact of substrate moisture content on growth and metabolic performance of black soldier fly larvae. Waste Management 127: 73-79.
Bertram, S.M. and Bellani, R., 2002. Influence of photoperiod on temporal mate signaling patterns in the Texas field cricket, Gryllus texensis. Journal of Insect Behavior 15: 607-615.
Bessa, L.W., Pieterse, E., Sigge, G. and Hoffman, L.C., 2017. Insects as human food; from farm to fork. Journal of the Science of Food and Agriculture 100: 5017-5022. https://doi.org/10.1002/jsfa.8860
Bjørge, J.D., Overgaard, J., Malte, H., Gianotten, N. and Heckmann, L.H., 2018. Role of temperature on growth and metabolic rate in the tenebrionid beetles Alphitobius diaperinus and Tenebrio molitor. Journal of Insect Physiology 107: 89-96.
Booth, D.T. and Kiddell, K., 2007. Temperature and the energetics of development in the house cricket (Acheta domesticus). Journal of Insect Physiology 53: 950-953.
Bosch, G., Oonincx, D.G.A.B., Jordan, H.R., Zhang, J., van Loon, J.J.A., van Huis, A. and Tomberlin, J.K., 2020. Standardisation of quantitative resource conversion studies with black soldier fly larvae. Journal of Insects as Food and Feed 6: 95-109.
Boulos, S., Tännler, A. and Nyström, L., 2020. Nitrogen-to-protein conversion factors for edible insects on the Swiss market: T. molitor, A. domesticus, and L. migratoria. Frontiers in Nutrition 7. https://doi.org/10.3389/fnut.2020.00089
Broeckx, L., Frooninckx, L., Slegers, L., Berrens, S., Noyens, I., Goossens, S., Verheyen, G., Wuyts, A. and Van Miert, S., 2021. Growth of black soldier fly larvae reared on organic side-streams. Sustainability 13: 12953. https://doi.org/10.3390/su132312953
Busvine, J.R., 1955. Simple methods for rearing the cricket (Gryllulus domesticus L.) with some observations on speed of development at different temperatures. Proceedings of the Royal Entomological Society of London. Series A, General Entomology 30: 15-18.
Cavazzana, E., Henrique, B., Leonardo, B., Borges, A., Carlos, T. and Martinez, B., 2007. Comparison of white-box and black-box models of a real hydraulic pumping system using a variable speeds drive. In: 19th international congress of mechanical engineering, November 5-9, 2007, Brası́lia, DF, Brazil. Available at: http://www.iau.dtu.dk/research/control/nnsysid.html
Chapman, R.F., 1998. The insects: structure and function. Cambridge University Press, Cambridge, UK, 483 pp.
Clifford, C.W., Roe, R.M. and Woodring, J.P., 1977. Rearing methods for obtaining house crickets, Acheta domesticus, of known age, sex, and instar. Annals of the Entomological Society of America 70: 69-74. https://doi.org/10.1093/aesa/70.1.69
Clifford, C.W. and Woodring, J.P., 1990. Methods for rearing the house cricket, Acheta domesticus (L.), along with baseline values for feeding rates, growth rates, development times, and blood composition. Journal of Applied Entomology 109: 1-14.
Collavo, A., Glew, R.H., Huang, Y.S., Chuang, L.T., Bosse, R. and Paoletti, M.G., 2005. House cricket small-scale farming. In: Paoletti, M.G. (ed.) Ecological implications of minilivestock: potential of insects, rodents, frogs and snails. Science Publishers, Enfield, NH, USA, pp. 515-540.
Cortes Ortiz, J.A., Ruiz, A.T., Morales-Ramos, J.A., Thomas, M., Rojas, M.G., Tomberlin, J.K., Yi, L., Han, R., Giroud, L. and Jullien, R.L., 2016. Insect mass production technologies. In: Dossey, T.A., Morales-Ramos, J.A. and Rojas, M.G. (eds.) Insects as sustainable food ingredients. Academic Press, London, UK, pp. 153-201.
Coudron, C.L., Deruytter, D. and Claeys, J., 2022a. The influence of wet feed pH on the growth of Tenebrio molitor larvae. Sustainability 14: 7841. https://doi.org/10.3390/su14137841
Coudron, C.L., Deruytter, D., Craeye, S. and Bleyaert, P., 2022b. Entomoponics: combining insect rearing and greenhouse vegetable production – a case study with Tenebrio molitor and high-wire cucumber cultivation. Journal of Insects as Food and Feed 8: 427-437.
Cymborowski, B., 1969. The effect of light on the circadian rhythm of locomotor activity of the house cricket (Acheta domesticus L.). Zoologica Poloniae 19: 85-95.
Cymborowski, B., 1973. Control of the circadian rhythm of locomotor activity in the house cricket. Journal of Insect Physiology 19: 1423-1440. https://doi.org/10.1016/0022-1910(73)90173-X
Deruytter, D., Coudron, C.L. and Claeys, J., 2021. The influence of wet feed distribution on the density, growth rate and growth variability of Tenebrio molitor. Journal of Insects as Food and Feed 7: 141-149.
Diener, S., Zurbrügg, C. and Tockner, K., 2009. Conversion of organic material by black soldier fly larvae: establishing optimal feeding rates. Waste Management and Research 27: 603-610.
Dobermann, D., Swift, J.A. and Field, L.M., 2017. Opportunities and hurdles of edible insects for food and feed. Nutrition Bulletin 42: 293-308.
Eilenberg, J., Vlak, J.M., Nielsen-LeRoux, C., Cappellozza, S. and Jensen, A.B., 2015. Diseases in insects produced for food and feed. Journal of Insects as Food and Feed 1: 87-102.
Fernandez-Cassi, X., Supeanu, A., Jansson, A., Boqvist, S. and Vagsholm, I., 2018. Novel foods: a risk profile for the house cricket (Acheta domesticus). EFSA Journal 16: e16082. https://doi.org/10.2903/j.efsa.2018.e16082
Finke, M.D., 2002. Complete nutrient composition of commercially raised invertebrates used as food for insectivores. Zoo Biology 21: 269-285.
Finke, M.D. and Oonincx, D., 2013. Insects as food for insectivores. In: Morales-Ramos, J.A., Guadalupe Rojas, M. and Shapiro-Ilan, D.I. (eds.) Mass production of beneficial organisms. Academic Press, Cambridge, MA, USA, pp. 583-616. https://doi.org/10.1016/B978-0-12-391453-8.00017-0
Fung, P.L., Zaidan, M.A., Timonen, H., Niemi, J.V., Kousa, A., Kuula, J., Luoma, K., Tarkoma, S., Petäjä, T., Kulmala, M. and Hussein, T., 2021. Evaluation of white-box versus black-box machine learning models in estimating ambient black carbon concentration. Journal of Aerosol Science 152: 105694. https://doi.org/10.1016/j.jaerosci.2020.105694
Ghouri, A.S.K. and Mcfarlane, J.E., 1958. Observations on the development of crickets. The Canadian Entomologist 90: 158-165. https://doi.org/10.4039/Ent90158-3
Halloran, A., Roos, N. and Hanboonsong, Y., 2017. Cricket farming as a livelihood strategy in Thailand. Geographical Journal 183: 112-124.
Han, X. and Heinonen, M., 2021. Development of ultra-high performance liquid chromatographic and fluorescent method for the analysis of insect chitin. Food Chemistry 334: 127577. https://doi.org/10.1016/j.foodchem.2020.127577
Hanboonsong, A. and Durst, P., 2020. Guidance on sustainable cricket farming – a practical manual for farmers and inspectors. FAO, Bangkok, Thailand, pp. 1-84. Available at: https://doi.org/https://doi.org/10.4060/cb2446en
Harsányi, E., Juhász, C., Kovács, E., Huzsvai, L., Pintér, R., Fekete, G., Varga, Z.I., Aleksza, L. and Gyuricza, C., 2020. Evaluation of organic wastes as substrates for rearing Zophobas morio, Tenebrio molitor, and Acheta domesticus larvae as alternative feed supplements. Insects 11: 1-18.
Inácio, A.C., Vågsholm, I., Jansson, A., Vaga, M., Boqvist, S. and Fraqueza, M.J., 2021. Impact of starvation on fat content and microbial load in edible crickets (Acheta domesticus). Journal of Insects as Food and Feed 7: 1143-1147.
IPIFF, 2021. EU novel food legislation. Available at: https://ipiff.org/insects-novel-food-eu-legislation-2/
Janssen, R.H., Vincken, J.P., van den Broek, L.A.M., Fogliano, V. and Lakemond, C.M.M., 2017. Nitrogen-to-protein conversion factors for three edible insects: Tenebrio molitor, Alphitobius diaperinus, and Hermetia illucens. Journal of Agricultural and Food Chemistry 65: 2275-2278.
Jensen, K., Kristensen, T.N., Heckmann, L.L. and Sørensen, J.G., 2017. Breeding and maintaining high-quality insects. In: van Huis, A. and Tomberlin, J.K. (eds.) Insects as food and feed: from production to consumption. Wageningen Academic Publishers, Wageningen, the Netherlands, pp. 174-198.
Jongema, Y., 2017. Worldwide list of recorded edible insects. Wageningen University & Research, Wageningen, the Netherlands. Available at: https://www.wur.nl/upload_mm/8/a/6/0fdfc700-3929-4a74-8b69-f02fd35a1696_Worldwide%20list%20of%20edible%20insects%202017.pdf
Kieruzel, M., 1976. The selection of shelter place by the house cricket. Acta Neurobiologiae Experimentalis 36: 561-580.
Kuo, C. and Fisher, B.L., 2022. A literature review of the use of weeds and agricultural and food industry by-products to feed farmed crickets (Insecta; Orthoptera; Gryllidae). Frontiers in Sustainable Food Systems 5. https://doi.org/10.3389/fsufs.2021.810421
Lecocq, A., Jensen, A.B. and Eilenberg, J., 2018. Diseases of insects in European production systems: diagnosis, prevention and management. Berliner und Münchener Tierärztliche Wochenschrift 131. https://doi.org/10.2376/0005-9366-18061
Loyola-Gonzalez, O., 2019. Black-box vs. white-box: understanding their advantages and weaknesses from a practical point of view. IEEE Access 7: 154096-154113.
Lundy, M.E. and Parrella, M.P., 2015. Crickets are not a free lunch: protein capture from scalable organic side-streams via high-density populations of Acheta domesticus. PLOS ONE 10: e0118785.
Magara, H.J.O., Niassy, S., Ayieko, M.A., Mukundamago, M., Egonyu, J.P., Tanga, C.M., Kimathi, E.K., Ongere, J.O., Fiaboe, K.K.M., Hugel, S., Orinda, M.A., Roos, N. and Ekesi, S., 2021. Edible crickets (Orthoptera) around the world: distribution, nutritional value, and other benefits – a review. Frontiers in Nutrition 7. https://doi.org/10.3389/fnut.2020.537915
Magara, H.J.O., Tanga, C.M., Ayieko, M.A., Hugel, S., Mohamed, S.A., Khamis, F.M., Salifu, D., Niassy, S., Sevgan, S., Fiaboe, K.K.M., Roos, N. and Ekesi, S., 2019. Performance of newly described native edible cricket Scapsipedus icipe (Orthoptera: Gryllidae) on various diets of relevance for farming. Journal of Economic Entomology 112: 653-664.
Mahavidanage, S., Fuciarelli, T.M., Xiaobing, L.I. and David Rollo, C., 2023. The effects of rearing density on growth, survival, and starvation resistance of the house cricket Acheta domesticus. Journal of Orthoptera Research 32: 25-31.
Marcucci, C., 2020. Food frontiers: insects as food, is the future already here? Mediterranean Journal of Nutrition and Metabolism 13: 43-52. https://doi.org/10.3233/MNM-190348
Marks, H.L., 1981. Role of water in regulating feed intake and feed efficiency of broilers. Poultry Science 60: 698-707.
McCluney, K.E. and Date, R.C., 2008. The effects of hydration on growth of the house cricket, Acheta domesticus. Journal of Insect Science 8: 32. https://doi.org/10.1673/031.008.3201
Morales-Ramos, J.A., Kelstrup, H.C., Guadalupe Rojas, M. and Emery, V., 2019. Body mass increase induced by eight years of artificial selection in the yellow mealworm (Coleoptera: Tenebrionidae) and life history trade-offs. Journal of Insect Science 19: 4. https://doi.org/10.1093/jisesa/iey110
Morales-Ramos, J.A., Rojas, M.G., Dossey, A.T. and Berhow, M., 2020. Self-selection of food ingredients and agricultural by-products by the house cricket, Acheta domesticus (Orthoptera: Gryllidae): a holistic approach to develop optimized diets. PLOS ONE 15: 1-30.
Mukherjee, S. and Mukherjee, A., 2022. Small-scale breeding and harvesting of house cricket Acheta domesticus. Acta Entomology and Zoology 3: 112-115. https://doi.org/10.33545/27080013.2022.v3.i2b.82
Murtaugh, M.P. and Denlinger, D.L., 1985. Physiological regulation of long-term oviposition in the house cricket, Acheta domesticus. Journal of Insect Physiology 31: 611-617. https://doi.org/10.1016/0022-1910(85)90059-9
Nakagaki, B.J. and Defoliart, A.R., 1991. Comparison of diets for mass-rearing Acheta domesticus (Orthoptera: Gryllidae) as a novelty food, and comparison of food conversion efficiency with values reported for livestock. Journal of Economic Entomology 84: 891-896.
Nijhout, H.F. and Wheeler, D.E., 1996. Growth models of complex allometries in holometabolous insects. The American Naturalist 148. https://doi.org/10.1086/285910
Nowosielski, J.W. and Patton, R.L., 1963. Studies on circadian rhythm of the house cricket, Gryllus domesticus L. Journal of Insect Physiology 9: 401-410.
Oloo, J.A., Ayieko, M. and Nyongesah, J.M., 2019. Acheta domesticus (cricket) feed resources among smallholder farmers in Lake Victoria region of Kenya. Food Science and Nutrition 8: 69-78.
O’Meara, F.M., Gardiner, G.E., O’Doherty, J.V. and Lawlor, P.G., 2020. Effect of water-to-feed ratio on feed disappearance, growth rate, feed efficiency, and carcass traits in growing-finishing pigs. Translational Animal Science 4: 630-640.
Oonincx, D.G.A.B. and Finke, M.D., 2021. Nutritional value of insects and ways to manipulate their composition. Journal of Insects as Food and Feed 7: 639-659.
Oonincx, D.G.A.B., Van Broekhoven, S., Van Huis, A. and Van Loon, J.J.A., 2015. Feed conversion, survival and development, and composition of four insect species on diets composed of food by-products. PLOS ONE 10: 1-20.
Oonincx, D.G.A.B., van Itterbeeck, J., Heetkamp, M.J.W., van den Brand, H., van Loon, J.J.A. and van Huis, A., 2010. An exploration on greenhouse gas and ammonia production by insect species suitable for animal or human consumption. PLOS ONE 5: 1-7.
Orinda, M.A., Mosi, R.O., Ayieko, M.A., Amimo, F.A. and Odinga, J.O., 2017. Growth performance of common house cricket (Acheta domesticus) and field cricket (Gryllus bimaculatus) crickets fed on agro-byproducts. Journal of Entomology and Zoology Studies 5: 1664-1668.
Patton, R.L., 1963. Rearing the house cricket, Acheta domesticus, on commercial feed. Annals of the Entomological Society of America 56: 250-251.
Patton, R.L., 1967. Oligidic diets for Acheta domesticus (Orthoptera: Gryllidae). Annals of the Entomological Society of America 60: 1238-1242.
Patton, R.L., 1978. Growth and development parameters for Acheta domesticus. Annals of the Entomological Society of America 71: 40-42.
Philippe, F.X. and Nicks, B., 2015. Review on greenhouse gas emissions from pig houses: production of carbon dioxide, methane and nitrous oxide by animals and manure. Agriculture, Ecosystems and Environment 199: 10-25. https://doi.org/10.1016/j.agee.2014.08.015
Quek, X.T., Liang, L., Tham, H.H., Yeo, H., Tan, M.K. and Tan, H.T.W., 2019. Are the growth and survival of Acheta domesticus comparable when reared on Okara, waste vegetables and premium animal feed? Journal of Insects as Food and Feed 6: 161-168.
Rehman, K., Rehman, A., Cai, M., Zheng, L., Xiao, X., Somroo, A.A., Wang, H., Li, W., Yu, Z. and Zhang, J., 2017. Conversion of mixtures of dairy manure and soybean curd residue by black soldier fly larvae (Hermetia illucens L.). Journal of Cleaner Production 154: 366-373.
Ritvanen, T., Pastell, H., Welling, A. and Raatikainen, M., 2020. The nitrogen-to-protein conversion factor of two cricket species – Acheta domesticus and Gryllus bimaculatus. Agricultural and Food Science 29: 1-5.
Roe, R.M., Clifford, C.W. and Woodring, J.P., 1980. The effect of temperature on feeding, growth, and metabolism during the last larval stadium of the female house cricket, Acheta domesticus. Journal of Insect Physiology 26: 639-644. https://doi.org/10.1016/0022-1910(80)90034-7
Rumbos, C.I., Adamaki-Sotiraki, C., Gourgouta, M., Karapanagiotidis, I.T., Asimaki, A., Mente, E. and Athanassiou, C.G., 2021. Strain matters: strain effect on the larval growth and performance of the yellow mealworm, Tenebrio molitor L. Journal of Insects as Food and Feed 7: 1195-1205.
Rumpold, B.A. and Schlüter, O.K., 2013. Nutritional composition and safety aspects of edible insects. Molecular Nutrition and Food Research 57: 802-823.
Rzeźnik, W. and Mielcarek, P., 2016. Greenhouse gases and ammonia emission factors from livestock buildings for pigs and dairy cows. Polish Journal of Environmental Studies 25: 1813-1821. https://doi.org/10.15244/pjoes/62489
Sandrock, C., Leupi, S., Wohlfahrt, J., Kaya, C., Heuel, M., Terranova, M., Blanckenhorn, W.U., Windisch, W., Kreuzer, M. and Leiber, F., 2022. Genotype-by-diet interactions for larval performance and body composition traits in the black soldier fly, Hermetia illucens. Insects 13: 424. https://doi.org/10.3390/insects13050424
Scriber, J.M. and Slansky, F., 1981. The nutritional ecology of immature insects. Annual Review of Entomology 26: 183-211.
Sorjonen, J.M., Valtonen, A., Hirvisalo, E., Karhapää, M., Lehtovaara, V.J., Lindgren, J., Marnila, P., Mooney, P., Mäki, M., Siljander-Rasi, H., Tapio, M., Tuiskula-Haavisto, M. and Roininen, H., 2019. The plant-based by-product diets for the mass-rearing of Acheta domesticus and Gryllus bimaculatus. PLOS ONE 14: 1-19.
Spranghers, T., Wouters, F., Schillewaert, S. and Frooninckx, L., 2021. Guide for insect farmers and processors towards sustainable and healthy insect rearing. Department of Agriculture and Fisheries, Schaarbeek, Belgium. Available at: https://publicaties.vlaanderen.be/view-file/46110
Sripontan, Y., Chiu, C.I., Tanansathaporn, S., Leasen, K., Manlong, K. and Papadopoulos, N., 2020. Modeling the growth of black soldier fly Hermetia illucens (Diptera: Stratiomyidae): an approach to evaluate diet quality. Journal of Economic Entomology 113: 742-751.
Tennis, P.S., Koonce, J.F. and Teraguchi, M., 1977. The effects of population density and food surface area on body weight of Acheta domesticus (L.) (Orthoptera: Gryllidae). Canadian Journal of Zoology 55: 2004-2010.
Tennis, P.S., Koonce, J.F. and Teraguchi, M., 1979. Studies on food size as a selection pressure on body size. I. effects of food size on fitness of two size strains of Acheta domesticus L. Evolution 33: 95-103.
Thompson, M., Owen, L., Wilkinson, K., Wood, R. and Damant, A., 2002. A comparison of the Kjeldahl and Dumas methods for the determination of protein in foods, using data from a proficiency testing scheme. Analyst 127: 1666-1668. https://doi.org/10.1039/B208973B
Vaga, M., Berggren, A. and Jansson, A., 2020a. Growth, survival and development of house crickets (Acheta domesticus) fed flowering plants. Journal of Insects as Food and Feed 7: 151-161.
Vaga, M., Berggren, A., Pauly, T. and Jansson, A., 2020b. Effect of red clover-only diets on house crickets (Acheta domesticus) growth and survival. Journal of Insects as Food and Feed 6: 179-189.
van Huis, A., 2013. Potential of insects as food and feed in assuring food security. Annual Review of Entomology 58: 563-583.
van Huis, A. and Oonincx, D.G.A.B., 2017. The environmental sustainability of insects as food and feed. A review. Agronomy for Sustainable Development 37: 43. https://doi.org/10.1007/s13593-017-0452-8
Van Huis, A., Van Itterbeeck, J., Klunder, H., Mertens, E., Halloran, A., Muir, G. and Vantomme, P., 2013. Edible insects. Future prospects for food and feed security. Food and Agriculture Organization of the United Nations, Rome, Italy. https://doi.org/10.1017/CBO9781107415324.004
Van Peer, M., Berrens, S., Noyens, I., Goossens, S., Coudron, C., Lau, T., Alvarez, C. and Van Miert, S., 2023. Testing side streams as substrate for insects. ValuSect. Available at: https://www.valusect.eu/node/64
Van Peer, M., Frooninckx, L., Coudron, C., Berrens, S., Álvarez, C., Deruytter, D., Verheyen, G. and Van Miert, S., 2021. Valorisation potential of using organic side streams as feed. Insects 12: 796. https://doi.org/10.3390/insects12090796
Waldbauer, G.P., 1968. The consumption and utilization of food by insects. Advances in Insect Physiology 5: 229-288.
Wang, Y., Liu, H. and Sun, Z., 2017. Lamarck rises from his grave: parental environment-induced epigenetic inheritance in model organisms and humans. Biological Reviews 92: 2084-2111.
Woodring, J.P., Clifford, C.W. and Beckman, B.R., 1979. Food utilization and metabolic efficiency in larval and adult house crickets. Journal of Insect Physiology 25: 903-912.
Zhou, F., Tomberlin, J.K., Zheng, L., Yu, Z. and Zhang, J., 2013. Developmental and waste reduction plasticity of three black soldier fly strains (Diptera: Stratiomyidae) raised on different livestock manures. Journal of Medical Entomology 50: 1224-1230.
