Microbial Dynamics during Industrial Rearing, Processing, andStorage of Tropical House Crickets (Gryllodes sigillatus) forHuman Consumption

Dries Vandeweyer,a Enya Wynants,a Sam Crauwels,b Christel Verreth,b Nikolaas Viaene,c Johan Claes,a Bart Lievens,b

Leen Van Campenhouta

aDepartment of Microbial and Molecular Systems (M2S), Lab4Food, KU Leuven, Geel Campus, Geel, BelgiumbDepartment of Microbial and Molecular Systems (M2S), Laboratory for Process Microbial Ecology andBioinspirational Management (PME & BIM), KU Leuven, De Nayer Campus, Sint-Katelijne-Waver, Belgium

cLittle Food CVBA, Brussels, Belgium

ABSTRACT In this study, the microbiota during industrial rearing, processing, and stor-age of the edible tropical house cricket, Gryllodes sigillatus, was investigated. To this end,we analyzed samples from the cricket feed, obtained before feeding as well as from thecages, and from the crickets during rearing, after harvest, and after processing into fro-zen, oven-dried, and smoked and oven-dried (smoked/dried) end products. Althoughthe feed contained lower microbial numbers than the crickets, both were dominated bythe same species-level operational taxonomic units, as determined by Illumina MiSeq se-quencing. They corresponded, among others, to members of Porphyromonadaceae, Fuso-bacterium, Parabacteroides, and Erwinia. The harvested crickets contained high microbialnumbers, but none of the investigated food pathogens Salmonella spp., Listeria monocy-togenes, Bacillus cereus, or coagulase-positive staphylococci. However, some possiblemycotoxin-producing fungi were isolated from the crickets. A postharvest heat treat-ment, shortly boiling the crickets, reduced microbial numbers, but an endospore load of2.4 log CFU/g remained. After processing, an increase in microbial counts was observedfor the dried and smoked/dried crickets. Additionally, in the smoked/dried crickets, ahigh abundance of a Bacillus sp. was observed. Considering the possible occurrence offood-pathogenic species from this genus, it is advised to apply a heat treatment whichis sufficient to eliminate spores. Nevertheless, the microbial numbers remained constantover a 6-month storage period, whether frozen (frozen end product) or at ambienttemperature (oven-dried and smoked/dried end products).

IMPORTANCE The need for sustainable protein sources has led to the emergence of anew food sector, producing and processing edible insects into foods. However, insightinto the microbial quality of this new food and into the microbial dynamics during rear-ing, processing, and storage of edible insects is still limited. Samples monitored for theirmicrobiota were obtained in this study from an industrial rearing and processing cycle.The results lead first to the identification of process steps which are critical for microbialfood safety. Second, they can be used in the construction of a Hazard Analysis and Criti-cal Control Points (HACCP) plan and of a Novel Food dossier, which is required inEurope for edible insects. Finally, they confirm the shelf-life period which was deter-mined by the rearer.

KEYWORDS food safety, Gryllodes sigillatus, high-throughput sequencing, insectrearing, metagenetics, microbial dynamics, shelf life

Edible insects are increasingly acknowledged as a sustainable alternative animalprotein source (1, 2). Consequently, a new food sector is being developed that

includes insect rearing companies and insect food producers. Other food-producing

Received 31 January 2018 Accepted 3 April2018

Accepted manuscript posted online 6 April2018

Citation Vandeweyer D, Wynants E, CrauwelsS, Verreth C, Viaene N, Claes J, Lievens B, VanCampenhout L. 2018. Microbial dynamicsduring industrial rearing, processing, andstorage of tropical house crickets (Gryllodessigillatus) for human consumption. ApplEnviron Microbiol 84:e00255-18. https://doi.org/10.1128/AEM.00255-18.

Editor Johanna Björkroth, University of Helsinki

Copyright © 2018 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Leen Van Campenhout,leen.vancampenhout@kuleuven.be.

D.V. and E.W. are co-first authors for this work.

FOOD MICROBIOLOGY

crossm

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 1Applied and Environmental Microbiology

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

sectors have already been subjected to extensive research regarding microbial foodsafety (3, 4) and can fall back on a clear legislative framework (regulation EC 2073/2005in Europe [5]). For the insect sector, in contrast, research on food safety and legislationare still in their infancy. It is already known from a few previous studies (6–10) thatduring an insect rearing process, substantial alterations in terms of microbial qualitycan occur over time. Furthermore, different edible insect species are being reared forhuman consumption. They can harbor a completely different microbial community (11,12), and hence, the microbial qualities of different insect species and productionprocesses should be investigated separately.

It has been suggested that insect feed is an important factor for the developmentof the insect microbial composition (7, 13–15). Furthermore, the feed may constitute apotential source of hazards of a microbiological origin, such as food pathogens andmycotoxins (7, 16, 17). In addition to the insect feed, the rearing environment and themanual practices of workers may also contribute to the microbiota of edible insects(18). In order to fully understand the relationship between these factors and themicrobial community composition, as well as possible food safety hazards, generalinsight into the microbial dynamics in the insect feed and the insects as such duringrearing is required. Furthermore, as already shown in previous research (19–22),postharvest processing of insects has a major impact on the microbial load. Processingtechniques, such as blanching, cooking, freezing, oven drying, and freeze-drying, arepresently applied for edible insects (2, 8, 19, 23, 24). Usually, companies rearing edibleinsects produce different end products from the same insect species, including frozen,dried, and seasoned insects. Many of these products are already introduced into themarket. Nevertheless, the impact of the processing steps on the microbial quality of theproducts has not yet been thoroughly assessed. Additionally, given their differentproduction processes, which likely result in different intrinsic properties per product,each product may differ in shelf life. Therefore, research is needed on the microbialstability of these different insect products.

The goal of this study was to investigate the microbial dynamics, including changesin microbial numbers and bacterial community composition, from industrial rearing toprocessing and storage of the tropical house cricket (Gryllodes sigillatus) reared forhuman consumption. For this purpose, the egg deposition medium, the feed beforeadministration (here referred to as “feed”), the feed present in the rearing cages andthus in contact with the crickets (here referred to as “substrate”), and the crickets weresampled and analyzed. Additionally, three end products, including crickets that werefrozen, oven-dried, and smoked and subsequently oven-dried (smoked/dried), were ana-lyzed after processing and packaging and during their 6-month shelf life, as proposedby the manufacturer (Little Food, Brussels, Belgium). To this end, samples were inves-tigated for their intrinsic parameters, microbial numbers, microbial community com-position (bacterial and fungal), and the presence of a selection of foodborne patho-gens.

RESULTSIntrinsic parameters. Water activity (aw), moisture content, and pH were deter-

mined for the egg deposition medium (“peat-peel mix”), the feed (days 0 and 26), thesubstrate (days 12, 26, and 37), homogenized insects (day 40), and frozen crickets. Dueto the small sample size, only water activity and moisture content were determined forthe dried crickets and the smoked/dried crickets. The peat-peel mix showed a high aw

and moisture content (average, 0.98 and 76.6%, respectively), whereas the pH averaged4.99 (Table 1). The feed consisted of a dry material with low aw (0.59 to 0.61) andmoisture content (10.3%). However, once administered to the crickets in the cage, thewater activity and moisture content of the substrate gradually increased to 0.78 and15.5%, respectively, at day 26. Toward the end of the rearing process (day 37), however,when the substrate was replenished more frequently, the water activity and themoisture content decreased again (P � 0.018 and 0.007, respectively [Table 1]). Thevalues for pH were not statistically different between the feed and substrate, irrespec-

Vandeweyer et al. Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 2

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

tive of the sampling days. After harvest, crickets were high in water activity andmoisture content (0.97 and 71.5% on average, respectively) and showed a near-neutralpH of 6.64 on average (Table 1).

Following heat treatment (bringing to a boil) of the crickets, the mean pH and moisturecontent significantly increased to 6.84 (P � 0.031) and 73.8% (P � 0.006), respectively. Nodifference was seen for water activity (Table 1). The frozen crickets did not show anydifference in intrinsic parameters from the heat-treated crickets (Table 2). The oven-driedand smoked/dried crickets were significantly lower in aw (P � 0.015 and 0.001,respectively) and moisture content (P � 0.000). During storage, aw (P � 0.024) andmoisture content (P � 0.018) of the smoked/dried crickets increased slightly (Table 2).

Plate counts. All samples collected during rearing (Fig. 1; also see Table S1 in thesupplemental material) were analyzed for their total viable count (TVC) and thenumbers of Enterobacteriaceae, lactic acid bacteria (LAB), aerobic bacterial endospores,and fungi. The peat-peel mix showed a high TVC of 8.5 log CFU/g, whereas other countsranged between 4.3 (LAB) and 6.9 (fungi) log CFU/g (Table 3). The feed, on the otherhand, contained lower numbers of microorganisms; the mean TVC ranged from 5.0 to5.4 log CFU/g and the other counts from 1.8 (LAB) to 4.5 (Enterobacteriaceae and

TABLE 1 Intrinsic properties during tropical house cricket rearinga

Sample Sample day

Intrinsic property

pH aw

Moisturecontent (%)

Peat-peel mix 0 4.99 � 1.09 0.98 � 0.00 76.6 � 3.9

Feed 0 5.51 � 0.08 a 0.59 � 0.02 a 10.3 � 0.4 a26 5.59 � 0.01 a 0.61 � 0.01 a 10.3 � 0.3 a

Substrate 12 5.57 � 0.11 a 0.73 � 0.02 a 13.0 � 1.0 a26 5.41 � 0.04 a 0.78 � 0.04 a 15.5 � 1.0 b37 5.49 � 0.06 a 0.61 � 0.04 b 9.8 � 0.2 c

Crickets 40 6.64 � 0.10 A 0.97 � 0.01 A 71.5 � 0.7 A

Heat-treated cricketsb 40 6.84 � 0.05 B 0.98 � 0.00 A 73.8 � 0.4 BaData are the mean � standard deviation values of three replicates. Means per product with the samelowercase letter within the same column do not differ significantly (P � 0.05). Means from differentproducts with the same uppercase letter within the same column do not differ significantly (P � 0.05).

bThe heat treatment consisted of bringing the crickets to a boil in a kettle with water.

TABLE 2 Intrinsic properties during storage of processed tropical house cricketsa

ProductStoragetime (mo)

Intrinsic property

pH aw

Moisturecontent (%)

Heat-treated cricketsb 0 6.84 � 0.05 A 0.98 � 0.00 A 73.8 � 0.4 A

Frozen crickets 0 6.85 � 0.06 aA 0.98 � 0.00 aA 73.6 � 0.6 aA3 6.95 � 0.04 a 0.97 � 0.01 a 74.3 � 1.0 a6 6.89 � 0.01 a 0.98 � 0.00 a 75.6 � 2.0 a

Oven-dried crickets 0 ND 0.35 � 0.08 aB 5.1 � 1.0 aB3 ND 0.36 � 0.01 a 4.4 � 0.4 a6 ND 0.36 � 0.01 a 6.0 � 0.3 a

Smoked and dried crickets 0 ND 0.24 � 0.03 aB 2.2 � 0.1 abB3 ND 0.30 � 0.03 ab 1.9 � 0.2 a6 ND 0.34 � 0.02 b 5.0 � 0.9 b

aData are the mean � standard deviation values of three replicates. Means per product with the samelowercase letter within the same column do not differ significantly (P � 0.05). Means from differentproducts with the same uppercase letter within the same column do not differ significantly (P � 0.05). ND,not determined.

bThe heat treatment consisted of bringing the crickets to a boil in a kettle with water.

Microbial Dynamics of a Cricket-Rearing Cycle Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 3

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

endospores) log CFU/g. The microbial load of the feed did not differ significantlybetween the samples taken at day 0 and day 26, except for the number of Enterobac-teriaceae, which was higher at day 0 (P � 0.017). Once the feed was added to the cages(becoming substrate), the mean TVC increased and ranged from 6.2 to 6.7 log CFU/g,but it did not differ significantly between sampling days 12, 26, and 37. Other averagecounts for the substrate ranged from 3.2 (fungi) to 5.0 (LAB) log CFU/g. Significantdecreases in microbial load were detected from day 12 to day 37 for Enterobacteriaceae(P � 0.018) and LAB (P � 0.023). During the rearing phase, the average TVCs for thecrickets ranged from 8.2 to 8.5 log CFU/g. The counts of Enterobacteriaceae ranged fromaverages of 7.2 to 7.5 log CFU/g, LAB from 6.7 to 7.8 log CFU/g, endospores from 3.5to 3.8 log CFU/g, and fungi from 5.4 to 6.0 log CFU/g. After heat treatment, all microbialcounts were reduced, with those for LAB and fungi being even below the detectionlimit (1 and 2 log CFU/g, respectively). The reduction was significant for all counts (P �

0.000, 0.018, 0.000, 0.002, and 0.004 for TVC, Enterobacteriaceae, LAB, endospores, andfungi, respectively).

Following processing, frozen crickets were stored for 6 months at �25 � 1°C, driedcrickets at 21 � 1°C, and smoked/dried crickets at 22 � 2°C. During processing and

FIG 1 (A) Schematic representation of the rearing and processing cycle of tropical house crickets. The three final end products aredepicted in green. The rearing period, from first instar to harvested adult cricket, took 40 days. In the manual harvesting step, Cricketswere harvested by shaking them out of the cardboard boxes into a circular plastic container. The crickets were then killed by submergingthem in hot water inside the container. The crickets were then rinsed thoroughly in a colander using running tap water for 5 min per batch.For the heat treatment step, the crickets were submerged in boiling water and until the water boiled again, which took 5 to 10 min. Thecrickets were salted after heat treatment by submerging them per batch in 4 liters of salted water (62.5 g/liter) for 40 min. Crickets weresmoked using a traditional beech wood smoker for 40 min at 80°C. In the drying step, crickets were spread out over a baking tray anddried overnight (10 h at 80°C). (B) Sampling plan throughout rearing, processing, and storage.

Vandeweyer et al. Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 4

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

storage of the crickets (Fig. 1), the TVC and numbers of Enterobacteriaceae and fungiwere determined (Table 4). After processing, the TVCs of both dried (P � 0.002) andsmoked/dried crickets (P � 0.000) were significantly higher than that of the heat-treated crickets. Additionally, the TVC of the smoked/dried crickets was higher than thatof the oven-dried crickets (P � 0.013). Enterobacteriaceae and fungi counts remainedbelow the detection limit, and the amount of endospores increased slightly but notsignificantly. During the 6-month storage period, all counts remained stable, except theTVC of the smoked/dried crickets, which slightly (but significantly, P � 0.009) decreasedover time (Table 4).

Identification of fungal isolates. Fungi isolated from the peat-peel mix (day 0), thefeed (days 0 and 26), the substrate at the end of the rearing period (day 37), and the cricketsafter harvest (day 40) were identified (Table S2). All isolates obtained from the peat-peelmix corresponded to the genus Trichoderma. For the feed, isolates corresponded to thegenera Aspergillus, Hyphopichia, Lichtheimia, and Penicillium. With regard to the sub-strate, isolates were identified as Aspergillus, Candida, Lichtheimia, Penicillium, andTrichoderma species. The isolates from the crickets corresponded to Aspergillus, Can-dida, Kodamaea, Lichtheimia, Tetrapisispora, Trichoderma, and Trichosporon species(Table S2).

TABLE 3 Microbial counts during tropical house cricket rearinga

Sample Sample day

Microbial count (log CFU/g)

Total viable count Enterobacteriaceae Lactic acid bacteria Endospores Fungi

Peat-peel mix 0 8.5 � 0.4 6.4 � 0.7 4.3 � 0.9 6.4 � 0.3 6.9 � 1.0

Feed 0 5.4 � 0.3 a 4.5 � 0.3 a 1.8 � 0.2 a 4.5 � 0.1 a 3.6 � 0.1 a26 5.0 � 0.3 a 3.6 � 0.2 b 1.8 � 0.2 a 4.4 � 0.3 a 3.7 � 0.4 a

Substrate 12 6.3 � 0.2 a 4.4 � 0.1 a 5.0 � 0.2 a 4.3 � 0.3 a 3.6 � 0.4 ab26 6.2 � 0.3 a 4.8 � 0.3 a 4.7 � 0.6 a 4.0 � 0.4 a 3.2 � 0.1 b37 6.7 � 0.3 a 3.6 � 0.2 b 3.6 � 0.5 b 4.8 � 0.5 a 4.0 � 0.3 a

Crickets 12 8.4 � 0.1 a 7.5 � 0.2 a 6.7 � 0.1 a 3.8 � 0.2 a 6.0 � 0.2 a26 8.2 � 0.1 a 7.3 � 0.5 a 7.2 � 0.1 ab 3.5 � 0.1 a 5.4 � 0.1 b40 8.5 � 0.2 aA 7.2 � 0.1 aA 7.8 � 0.4 bA 3.7 � 0.3 aA 5.6 � 0.3 abA

Heat-treated cricketsb 40 2.6 � 0.5 B �1.5 � 0.9 B �1.0 � 0.0 B 2.4 � 0.4 B �2.0 � 0.0 BaData are the mean � standard deviation values of three replicates. Means per sample with the same lowercase letter within the same column do not differsignificantly (P � 0.05). Means from harvested crickets and heat-treated crickets (day 40) with the same uppercase letter within the same column do not differsignificantly (P � 0.05).

bThe heat treatment consisted of bringing the crickets to a boil in a kettle with water.

TABLE 4 Microbial counts during storage of processed tropical house cricketsa

ProductStoragetime (mo)

Microbial count (log CFU/g)

Total viable count Enterobacteriaceae Endospores Fungi

Heat-treated cricketsb 0 2.6 � 0.5 A �1.5 � 0.9 A 2.4 � 0.4 AB �2.0 � 0.0 aA

Frozen crickets 0 2.4 � 0.4 aA �1.0 � 0.0 aA 2.0 � 0.4 aA �2.0 � 0.0 aA3 2.5 � 0.3 a �1.1 � 0.1 a 2.4 � 0.4 a �2.1 � 0.1 a6 2.2 � 0.1 a �1.0 � 0.0 a 2.2 � 0.0 a �2.0 � 0.0 a

Oven-dried crickets 0 4.3 � 0.0 aB �1.0 � 0.0 aA 2.4 � 0.4 aAB �2.0 � 0.0 aA3 3.9 � 0.8 a �1.0 � 0.0 a 2.3 � 0.5 a �2.0 � 0.0 a6 3.9 � 0.8 a �1.1 � 0.1 a 2.5 � 0.5 a �2.1 � 0.1 a

Smoked and dried crickets 0 7.9 � 0.1 aC �1.0 � 0.0 aA 3.4 � 0.6 aB �2.0 � 0.0 aA3 7.4 � 0.3 ab �1.0 � 0.0 a 3.9 � 0.6 a �2.0 � 0.0 a6 7.0 � 0.2 b �1.0 � 0.0 a 3.2 � 0.0 a �2.0 � 0.0 a

aData are the mean � standard deviation values of three replicates. Means per product with the same lowercase letter within the same column do not differsignificantly (P � 0.05). Means from unstored (0 months) products with the same uppercase letter within the same column do not differ significantly (P � 0.05).

bThe heat treatment consisted of bringing the crickets to a boil in a kettle with water.

Microbial Dynamics of a Cricket-Rearing Cycle Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 5

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

Pathogen detection. At the end of the rearing cycle, harvested crickets (day 40) andthe substrate (day 37) were assessed for the presence of a number of foodbornepathogenic bacteria. Neither Salmonella spp. nor Listeria monocytogenes were detectedin the samples (absent in 25 g). Additionally, Bacillus cereus and coagulase-positivestaphylococcal counts were below the detection limit of 100 CFU/g.

Culture-independent analyses. A selection of samples (Table S1), including sam-ples from the peat-peel mix (day 0), the feed (day 0), the substrate (days 12 and 26),crickets during the rearing phase (days 12 and 26), and crickets after harvest andprocessing (Fig. 1), were subjected to high-throughput amplicon-based sequencing.Relative abundances and diversity indices were averaged over all DNA extracts ofreplicate samples. The average coverages, based on the Chao1 estimator, ranged from96.5% to 99.1%, indicating that the majority of the bacterial community members wererecovered (Table 5). Indices for species richness (observed richness and Chao1 [25])showed that the feed and substrate contained most bacterial species, while the leastdiversity was observed in the peat-peel mix, although its mean Shannon index (26) waswithin the range of the other samples (Table 5).

Identification of the operational taxonomic units (OTUs) (Data Set S3 and Table S4)revealed that the most abundant phyla in the peat-peel mix and feed were Proteobacteria(47.7% � 11.9% and 39.6% � 13.0%, respectively) and Bacteroidetes (38.0% � 13.7% and43.3% � 10.2%, respectively). In the feed, the Firmicutes was also among the mostabundant phyla (12.6% � 0.6%). Once the feed had been administered inside the cage,Firmicutes became more abundant (17.7% � 2.6% at day 12 and 20.2% � 1.7% at day 26)alongside Bacteroidetes and Proteobacteria (65.8% � 5.6% and 14.1% � 2.7%, respectively,at day 12; 47.4% � 9.9% and 29.2% � 11.7%, respectively, at day 26). During rearing andprocessing, Bacteroidetes remained the most abundant phylum in the crickets (rangingfrom 35.8% � 0.2% in smoked/dried crickets to 71.2% � 1.7% in the frozen crickets),followed by Firmicutes (ranging from 5.4% � 0.3% in the frozen crickets to 56.6% � 1.9%in the smoked/dried crickets) and Proteobacteria (ranging from 4.9% � 1.4% in thesmoked/dried crickets to 16.0% � 2.5% in the heat-treated crickets). Other phyla werepresent in abundances below 10% in any sample of the data set.

A total of 337 OTUs were identified throughout the data set (Data Set S3). Thepeat-peel mix (Fig. 2) was dominated (average OTU abundance of �5% in any sample)by OTUs corresponding to members of Chitinophagaceae (OTUs 15, 19, and 36), aChryseobacterium sp. (OTU 22), a Dyella sp. (OTU 23), a Burkholderia sp. (OTU 28), anda Rhodanobacter sp. (OTU 44). The feed and substrate samples (Fig. 2) were all

TABLE 5 Diversity indices for samples subjected to metagenetic analysis in this studya

Product Sampling moment

Diversity index

Observed richness Chao1b Coverage (%)c Shannon indexd

Peat-peel mix Start of rearing 90 � 5 92.70 � 7.99 96.74 � 3.45 3.22 � 0.20

Feed Start of rearing 128 � 12 131.30 � 7.92 97.01 � 3.30 3.45 � 0.28

Substrate Rearing day 12 127 � 12 A 128.28 � 12.44 A 99.00 � 0.56 A 3.39 � 0.22 ARearing day 26 130 � 4 A 131.75 � 4.76 A 98.31 � 1.24 A 3.59 � 0.02 A

Crickets Rearing day 12 115 � 7 ABC 116.33 � 7.14 ABC 99.08 � 0.33 A 3.45 � 0.02 ARearing day 26 120 � 1 B 122.53 � 0.83 B 97.94 � 0.58 A 3.48 � 0.06 AEnd of rearing 115 � 11 ABC 117.05 � 8.27 ABCD 97.75 � 2.15 A 3.53 � 0.01 AAfter heat treatment 101 � 4 AC 104.43 � 4.77 CD 96.52 � 1.47 A 3.18 � 0.02 BAfter freezing 104 � 2 A 106.05 � 1.41 ACD 97.83 � 1.13 A 3.16 � 0.03 BAfter oven drying 114 � 5 AB 116.35 � 4.11 AB 98.19 � 0.71 A 3.20 � 0.18 BAfter smoking 94 � 1 C 96.10 � 0.14 D 97.82 � 1.62 A 2.66 � 0.05 C

aSequences were grouped into OTUs defined by 97% sequence identity at the 16S rRNA gene (V4 region, 250 bp). Data are the mean � standard deviation values of twoanalyzed DNA extracts from two replicates per sampling moment. Means per product with the same letter within the same column do not differ significantly (P � 0.05).

bChao1 richness estimator is the total number of OTUs estimated by infinite sampling. A higher number indicates a higher richness (25).cCoverage was determined by using the equation (observed richness/Chao1 estimate) � 100.dShannon-Wiener diversity index is the index to characterize species diversity based on species richness as well as their relative abundance. A higher value representsmore diversity (26).

Vandeweyer et al. Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 6

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

dominated by the same group of OTUs, consisting of members of the Porphyromon-adaceae family (OTUs 1, 2, and 4), an Erwinia sp. (OTU 10), a Rhizobiales sp. (OTU 17), anda Pseudomonas sp. (OTU 49). More than 40% of the sequences recovered belonged toOTUs represented by �3% of the sequences in all samples. Both processed andunprocessed cricket samples (Fig. 3) were abundant in OTUs corresponding to mem-bers of Porphyromonadaceae (OTUs 2 and 4) and Enterobacteriaceae (OTU 8), a Bacte-roides sp. (OTU 3), a Fusobacterium sp. (OTU 5), a Parabacteroides sp. (OTU 6), and anErwinia sp. (OTU 10). During rearing, OTU 1 (a Porphyromonadaceae sp.) decreased inaverage relative abundance from 5% to below 1%. After harvest, crickets showed a high

FIG 2 Relative abundance (%) of OTUs present in the peat-peel mix, feed and substrate. Data are mean values oftwo extracts per sample from two samples. Error bars represent the standard deviation. Only OTUs represented byan average relative abundance of more than 3% of sequences in any sample are shown. OTUs with a mean relativeabundance of less than 3% are grouped in “Other OTUs (�3%)”.

FIG 3 Relative abundance (%) of OTUs present in the crickets during the rearing phase and after processing. Data are mean values of two extracts per samplefrom one (crickets at day 40 and smoked/dried crickets) or two cricket samples. Error bars represent the standard deviation. Only OTUs represented by anaverage relative abundance of more than 3% of sequences in any sample are shown. OTUs with a mean relative abundance of less than 3% are grouped in“Other OTUs (�3%)”.

Microbial Dynamics of a Cricket-Rearing Cycle Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 7

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

abundance of two Acinetobacter spp. (OTUs 34 and 52), and smoked/dried cricketsshowed a large abundance (�40%) of a Bacillus species (OTU 9). Also here, a largepercentage of OTUs showed a relative abundance of below 3% in any sample.

DISCUSSION

The goal of the present study was to achieve more insight into the microbialdynamics during rearing, processing, and storage of tropical house crickets sold forhuman consumption. To this end, crickets during the rearing phase and after process-ing into three end products, as well as the feed and substrate samples, were assessedfor their microbial quality. Additionally, for the three end products, microbial countswere monitored during the shelf life proposed by the manufacturer (Little Food), i.e., 6months.

Preharvest. The peat-peel mix contained high numbers of microorganisms, ofwhich the most abundant were typical soilborne bacteria, such as Dyella (OTU 23),Burkholderia (OTU 28), and Rhodanobacter (OTU 44) species and members of Chitin-ophagaceae (OTU 36) and Rhizobiales (OTU 17) (27, 28). In contrast, none of these OTUswere recovered in substantial abundance in the other samples. This suggests that thehatchlings do not transfer these bacteria to the rearing cages, nor that they retain thesebacteria within their gastrointestinal tract. The feed, on the other hand, contained lowermicrobial numbers, which greatly increased after the feed was placed inside the cage.That effect may be attributed to the increase in water activity and moisture content dueto absorption from the environment, thereby facilitating microbial growth and/orcross-contamination from contact with crickets and possible defecation into the sub-strate. As the crickets grew, the substrate was replenished more frequently toward theend of the rearing cycle to provide them with sufficient resources. This “dilution” maynot only explain the decrease in water activity and moisture content toward day 37 inthe substrate but also the observed decrease in the number of lactic acid bacteria andEnterobacteriaceae. The microbial numbers of the substrate were consistently lowerthan those of the crickets for all counts, except for the endospores, suggesting that thecrickets are a better matrix for microbial growth than the substrate. This is in contrastto the results obtained for the rearing of lesser mealworms (Alphitobius diaperinus) (8),where the larvae are reared within the substrate, which also contains their excretedfeces and exuviae.

Even though the microbial numbers of the feed and substrate are generally lowerthan those of the crickets, a striking similarity was observed in community compositionbetween the feed, the substrate, and the crickets. Indeed, for example, OTUs corre-sponding to Porphyromonadaceae (OTUs 1, 2 and 4), a Fusobacterium sp. (OTU 5), aParabacteroides sp. (OTU 6), and an Erwinia sp. (OTU 10) were abundant in all samples.The OTU most abundant in the feed belonged to the genus Erwinia (OTU 10). Mostmembers of this genus are plant-associated and plant-pathogenic bacteria (28), whichcould explain their high relative abundances in the plant-based feed. These resultssuggest that the feed is an important source for the development of the cricketmicrobiota. Similar results were obtained during a previous study on lesser mealworms,where 50 out of 70 bacterial OTUs identified from the feed were also detected in thelarvae (8). In that study, however, the microbial diversity of the larvae decreased duringrearing, with 22 OTUs obtained for the harvested larvae. In the present study, thecricket microbial community composition was highly diverse and did not significantlychange during rearing, as indicated by the diversity indices as well as by the largeportion of OTUs represented by �3% of the sequences. This observation was alsoreported by Vandeweyer et al. (11), where both the diversity indices and the numberof OTUs represented by �3% were higher for crickets than the values observed formealworms. Crickets thus harbor a bacterial community that is remarkably morecomplex than that of mealworms, both during and at the end of their rearing period,and with more dominating organisms. Also with regard to community composition,similarities were observed between the crickets in the present study and those ana-lyzed by Vandeweyer et al. (11). Indeed, those authors also report the presence of OTUs

Vandeweyer et al. Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 8

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

corresponding to the families Enterobacteriaceae and Pseudomonadaceae and thegenera Bacteroides, Parabacteroides, Erwinia, and Fusobacterium in fresh crickets. Alto-gether, it is reasonable to assume that Bacteroides spp. and Parabacteroides spp.,among other Porphyromonadaceae spp., are typical members of the endogenousintestinal bacterial community of crickets. Also, a Fusobacterium sp. has already beenobserved to be a member of the tropical house cricket microbiome (11). Noteworthy isthe presence of Acinetobacter spp. in our data set, e.g., OTUs 34 and 52 in the samplefrom harvested crickets. Acinetobacter species are widely distributed in nature andcommonly occur in soil and water but also in insect guts and plant-related environ-ments. Furthermore, the genus Acinetobacter contains multiple nosocomial opportu-nistic pathogens, of which A. baumannii is the most well-known (29). Interestingly,Acinetobacter species were previously found in carrots (30) and their wash water (31),which may explain their appearance in the harvested crickets, since exclusively carrotswere administered during the final days before harvest.

With regard to the microbial quality of fresh harvested crickets, the plate countnumbers obtained in our study were comparable to those obtained by Vandeweyer etal. (32) for tropical house crickets and house crickets (Acheta domesticus). Since thebeginning of the rearing process, most microbial counts remained stable within thecrickets, except for the LAB, which showed a significant increase in numbers over time.This significant rise may be explained by the good growth of (a few) LAB species whichmay be well adapted to the cricket gut environment. The retrieved Enterococcus sp.(OTU 21) in this study is a possible candidate for this hypothesis, because the abun-dance of this OTU in the crickets during the rearing phase rose since day 12. Moreover,Enterococcus species have been frequently detected in other fresh edible cricketsamples (11).

None of the four investigated foodborne pathogens were recovered from thecrickets after harvest or from the substrate, suggesting they were not present in therearing cycle under study. To our knowledge, L. monocytogenes has not yet beendetected in edible insects for human consumption (8, 32–35). However, Salmonellaspp., B. cereus, and Staphylococcus aureus have been reported in products, includingrhinoceros beetles (Oryctes monoceros), grasshoppers, mealworms (Tenebrio molitor),and/or house crickets (A. domesticus) (34–37). Of note, it remains to be investigatedwhether a pathogen possibly present in the insect feed may contaminate the insect (38,39) and thus pose a hazard for food safety of the end product. Although no foodpathogens were detected, some possible mycotoxin-forming fungi, more specificallyAspergillus spp. and Penicillium spp., were recovered from the feed, substrate, and/orcrickets. Future research should focus on the presence of these fungal genera in insectsand their potential to produce mycotoxins. Mycotoxins can be very heat resistant (40)and, if present, will likely not be reduced by heat treatment and further processing andwill be able to cause mycotoxicosis. Fungal infections (mycoses) caused by, e.g.,Aspergillus or Candida spp., via consumption of the crickets, on the other hand, arehighly unlikely, as the heat treatment was shown to reduce fungi to below thedetection limit (�2 log CFU/g).

Postharvest. The application of heat caused all counts to drop significantly. How-ever, a substantial number of endospores remained, as was expected based on previ-ous research on (lesser) mealworms and crickets (8, 19, 21). The unexpected increase ofbacterial counts after processing the crickets into oven-dried and smoked/dried endproducts can likely be explained by several factors, including cross-contaminationthrough the equipment and installations used, postcontamination through humaninteraction while removing legs and during packaging, and/or, for smoked/driedcrickets, the possibility of microbial growth during subsequent processing steps, in-cluding freezing and thawing cycles. To reduce risks for such contamination, it isadvised to incorporate good hygiene and manufacturing practices, such as the wearingof gloves and proper cleaning and disinfection of equipment. During the 6-monthstorage of the products, only the water activity and the moisture content of the

Microbial Dynamics of a Cricket-Rearing Cycle Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 9

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

smoked/dried crickets changed significantly, which indicates that the packaging tech-nique used (glass tube with cork stop) allows moisture to enter the product. The aw

value, however, never rose above 0.60 to allow microbial growth (41) and thereforeexplains the stable microbial quality during the proposed shelf life.

Heat-treated crickets had a bacterial community composition similar to that of thecrickets during the rearing phase. However, it should be noted that the heat treatment,although reducing microbial numbers significantly, possibly did not break down allbacterial DNA, which can explain the comparable relative abundances of the recoveredOTUs. Many genera observed in this study were also encountered in previous researchon processed crickets performed by Garofalo et al. (12). Highly remarkable, however, isthe appearance of a strongly abundant Bacillus species (OTU 9, 40.9%), a typicalspore-forming bacterium, in the smoked/dried crickets. The succession of processingsteps to obtain the smoked/dried crickets may have triggered subsequent cycles ofspore formation and germination, while the relative abundance of the backgroundmicrobiota was reduced. The high abundance of this OTU also explains the significantlylower bacterial diversity in the smoked/dried crickets. As the genus Bacillus contains thefood pathogen B. cereus, special attention should be paid to the heat treatment, eventhough B. cereus was not detected using the classical enumeration method (ISO7932:2004 [42]) performed on the harvested crickets. It is advised to apply a heattreatment which is sufficient to eliminate endospores and to decrease the amount offreeze-thaw cycles in the production process of such insect products to a minimum.

Conclusions. In this study, the microbial dynamics during rearing, processing, andstorage of tropical house crickets were investigated. The most abundant bacterialcommunity members identified from the cricket feed (e.g., Porphyromonadaceae spp.,a Bacteroides sp., a Parabacteroides sp., an Erwinia sp., and a Fusobacterium sp.) werealso recovered from cricket samples, suggesting the feed to be an important source forthe cricket microbiota. High microbial numbers of crickets during the rearing phasewere significantly reduced by a heat treatment. Neither Salmonella spp., Listeria mono-cytogenes, coagulase-positive staphylococci, nor Bacillus cereus were recovered fromthe crickets, but some mycotoxin-producing fungal genera were isolated. Furtherresearch on the possible presence of mycotoxins in insects is therefore advised. Afterprocessing into dried and smoked/dried end products, an increase in total microbialcount could be detected as a result of postcontamination, but the microbial communitycomposition remained comparable to that of the live crickets. However, a high abun-dance of a Bacillus sp. was introduced in the smoked/dried crickets. It is advised toapply a heat treatment sufficient to eliminate endospores and to minimize the numberof freeze-thawing cycles during processing. During their 6-month shelf life, the micro-bial quality of the cricket products, as indicated by their microbial numbers, remainedstable. The shelf life recommended by the manufacturer (Little Food) is thereforeacceptable from a microbiological point of view.

MATERIALS AND METHODSIndustrial rearing cycle. A complete rearing cycle in a Belgian company rearing crickets for human

consumption (Little Food, Brussels, Belgium) was monitored. An overview of the whole industrial cricketproduction process, including postharvest treatments, is given in Fig. 1A. Eggs were laid by adult cricketsin the final 1 to 2 days before harvest into a mixture of peat soil and coconut peel (peat-peel mix) in small(17.5 by 13 by 5 cm) plastic containers. These plastic containers were then placed in a larger containeron top of a pile of egg cardboard where the freshly emerged nymphs could reside. The nymphs wereremoved from the container and placed in a larger cage (approximately 2 by 1 by 1 m), consisting of awooden skeleton and Perspex walls, which was open on the upper side. Inside the cage, egg cartonswere piled up to create a dark habitat with crevices. All cages were situated in a ventilated room at anaverage temperature of 31°C and an average relative humidity of 70%. Artificial light was present onaverage 8 h/day except during weekends. A specialized cricket feed (main components were wheat bran,linseed flakes, and sunflower seed flakes) was added on a cardboard plate on top of the egg cartons, andwater was presented separately in a plastic dispenser. New feed was added one to three times a weekin quantities ranging from 1 to 2 kg, with both frequency and quantity increasing as the crickets aged.The water bowl was refilled when empty. Two days before harvest, only carrots were provided as feedand water source and, according to the rearing company, to improve reproduction and taste. Addition-ally, a small plastic container with peat and coconut peel (see above) was placed inside the cage foroviposition. After 40 days, the crickets were harvested, and the cages were cleaned with a brush and, if

Vandeweyer et al. Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 10

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

necessary, with water containing disinfectant. Subsequently, the crickets were killed by submersion inhot water (60°C) and rinsed (5 min) with regular tap water. Afterwards, they were given a heat treatmentby placing them in a kettle with boiling water and keeping them submerged until the water boiled again(after 5 to 10 min). Finally, heat-treated crickets were further processed into three end products. Cricketswere either frozen to �20°C (frozen crickets), oven-dried overnight at 80°C (dried crickets), or smoked.The smoking process involved a combination of salting (submerging in brine of 62.5 g NaCl/liter for 40min), freezing to �20°C (crickets were stored in the freezer until they were smoked), thawing, smoking(traditional beech wood smoker for 40 min at 80°C), and finally oven-drying overnight at 80°C (smoked/dried crickets).

Sample collection. During the rearing cycle, samples were taken from the peat-peel mix, the feedbefore addition, the feed in the cage (substrate), and the crickets every 2 weeks during the rearing cycle(Fig. 1B). At every sampling time point, three replicates were obtained per sample (i.e., 3-fold sampling).For substrates and crickets, each replicate was collected from a separate cage. At the end of the cycle,crickets were sampled after harvest and after heat treatment, again in triplicate. After processing, ninepacked samples of all three end products (frozen, dried, and smoked/dried crickets) were obtained aswell.

Storage conditions. All samples taken during and at the end of the rearing cycle (fresh samples) aswell as the heat-treated samples were kept at 3°C for a maximum of 24 h (cricket samples) or 48 h (feedsamples) until analyses. Of the end products, three packages were analyzed immediately after sampling,while the remaining samples were saved for long-term storage and evaluated in 3-fold after 3 and 6months of storage (Fig. 1B). Frozen cricket samples were stored in sealed plastic bags at �25°C. Dried andsmoked/dried cricket samples were stored in individual glass tubes with a cork stop (which is thepackaging used by the company to commercialize the crickets) at ambient temperature. During theentire storage period, temperatures were monitored using data loggers (Escort iLog internal sensor; VWRInternational, Leuven, Belgium).

Intrinsic parameters. All selected cricket samples (Table S1) were pulverized prior to analyses, asdescribed by Stoops et al. (43). Samples of the frozen end products were thawed for 4 h at 3°C beforepulverization. Peat-peel mix, feed, and substrate samples were analyzed without preparation. Wateractivity was measured using a water activity meter (LabMaster aw; Novasina, Lachen, Switzerland) untilthe aw and temperature (25°C) were stable for 15 and 5 min, respectively. The moisture content wasdetermined by calculating the difference in weight of 3 g (dried samples) or 5 g (other samples) of theinitial sample before and after oven drying for 17 h at 105°C. The pH was measured directly in the(homogenized) samples using a digital pH meter (Portamess 911; Knick, Berlin, Germany, with SI analyticselectrode; Mainz, Germany).

Microbiological plate counts. For the selected (pulverized) samples (Table S1), plate counts wereperformed according to the ISO standards for microbial analyses of food and feed as compiled by Dijket al. (44). TVCs were determined on plate count agar (PCA; Biokar Diagnostics, Beauvais, France) afterincubation at 30°C for 72 h. The number of LAB was determined on de Man-Rogosa-Sharpe (MRS) agar(Biokar Diagnostics), with the addition of sorbic acid (0.14%, causing the pH to drop to 5.20) to preventfungal growth, after incubation for 72 h at 30°C. Enterobacteriaceae were determined on violet red bileglucose (VRBG) medium (Biokar Diagnostics) after incubation at 37°C for 24 h. Aerobic bacterialendospores were determined by subjecting the 10�1 dilution to a heat shock treatment (10 min at 80°C),followed by a 10-fold serial dilution, plating onto PCA, and incubation at 37°C for 48 h. Fungi weredetermined on dichloran rose bengal chloramphenicol (DRBC) medium (Biokar Diagnostics) after incu-bation at 25°C for 6 days. The relative humidity of the atmosphere during incubation was 54%.

Identification of fungal isolates. For the feed (day 0), the peat-peel mix (day 0), the substrate (day37), and the crickets (day 40), a selection of fungal colonies with distinct morphology was picked fromthe DRBC medium for further identification (if possible, 10 colonies per sample type). Subsequently,isolates were grown on potato dextrose agar (PDA; Biokar Diagnostics) and incubated at 25°C. After 2 to7 days of incubation (depending on the growth), genomic DNA was extracted from purified strains usingthe phenol-chloroform DNA extraction procedure described by Lievens et al. (45). Identifications wereperformed by amplifying and sequencing the internal transcribed spacer (ITS) region (ITS1-5.8S ribo-somal DNA [rDNA]-ITS2) as described previously (45). The obtained sequences were compared with thenucleotide database in GenBank (46) (excluding unclassified and environmental entries), and isolateswere assigned to the highest taxonomic rank possible.

Pathogen detection. For three replicate cricket and substrate samples at the end of the rearingphase (day 40 and day 37, respectively), the presence of four food pathogens was assessed. The presenceof Salmonella was assessed according to ISO 6579-1:2017 (47) (absence in 25 g) and the presence of L.monocytogenes according to AFNOR BRD 07/4-09/98 (48) (absence in 25 g). Furthermore, detection of B.cereus was performed according to ISO 7932:2004 (42) (plate count) and the prevalence of coagulase-positive staphylococci according to AFNOR 3M 01/9-04/03 B (49) (plate count).

16S rRNA gene amplicon sequencing. The bacterial community composition of the selectedsamples (Table S1) was determined using Illumina MiSeq sequencing of partial 16S rRNA gene amplicons(V4 region, 250 bp). To this end, two replicates of each sample were pulverized as described for theintrinsic property and plate count analyses. Subsequently, DNA extraction, PCR amplification (primerdesign shown in Table S5), library preparation, sequencing, sequence processing, and diversity analyseswere performed as described by Wynants et al. (8). For each pulverized replicate, genomic DNA wasextracted in duplicate, resulting in a total of 4 DNA extracts per sample. Downstream diversity analysesused data rarefied to 1,700 sequences per DNA extract. For the harvested (fresh) and the smoked/driedcrickets, only two DNA extractions (of one replicate) delivered useful sequences; the others were not

Microbial Dynamics of a Cricket-Rearing Cycle Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 11

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

retained for data analysis. Sequences were clustered into operational taxonomic units (OTUs) based ona 97% similarity cutoff as proxies for species. The taxonomic origin of each OTU was determined to thegenus level with the SINTAX algorithm implemented in USEARCH (50) based on the Silva Living TreeProject (LTP) version 123 database. Taxonomic assignments were considered reliable when bootstrapconfidence values exceeded 0.80 (Data Set S3). In case the genus could not be determined reliably(bootstrap value, �0.80) based on the Silva database, OTU representative sequences were compared tothe nucleotide database in GenBank (excluding uncultured/environmental entries; Table S4). Chao1 andShannon-Wiener diversity indices were calculated using the R package Phyloseq (version 1.19.0) (51).

Statistical analyses. Differences in the intrinsic parameters, microbial counts, and diversity param-eters (OTU richness, Chao1, coverage, and Shannon-Wiener indices) during rearing, processing, andstorage of the crickets were analyzed by one-way analysis of variance (ANOVA), followed by Tukey’s posthoc test. In case of unequal variances, Welch’s ANOVA with a Games-Howell post hoc test was used. Alltests were performed with SPSS Statistics 23 (IBM, New York, NY, USA) and considered significant at a Pvalue below 0.05.

Accession number(s). The obtained sequences from the identified fungal isolates have beendeposited in GenBank under the accession numbers MG655272 to MG655305. The sequences obtainedfrom the Illumina MiSeq platform were deposited in the Sequence Read Archive (SAMN08032682 toSAMN08032721) under BioProject accession PRJNA418072. Additionally, representative sequences perOTU have been submitted to GenBank under accession numbers MG558004 to MG558332.

SUPPLEMENTAL MATERIAL

Supplemental material for this article may be found at https://doi.org/10.1128/AEM.00255-18.

SUPPLEMENTAL FILE 1, PDF file, 4.0 MB.SUPPLEMENTAL FILE 2, XLSX file, 0.1 MB.

ACKNOWLEDGMENTSThis work was supported by Flanders Innovation & Entrepreneurship (VLAIO, project

141129) and the Belgian Federal Public Service Health, Food Chain Safety and Envi-ronment (project EDINCO, RT 15/9).

We declare no conflicts of interest in the collaboration with the insect-rearingcompany Little Food.

REFERENCES1. Oonincx DGAB, de Boer IJM. 2012. Environmental impact of the

production of mealworms as a protein source for humans—a lifecycle assessment. PLoS One 7:e51145. https://doi.org/10.1371/journal.pone.0051145.

2. van Huis A, Van Itterbeeck J, Klunder HC, Mertens E, Halloran A, Muir G,Vantomme P. 2013. Edible insects: future prospects for food and feedsecurity. Food and Agricultural Organization of the United Nations,Rome, Italy.

3. Adams MR, Moss MO, McClure P. 2015. Food microbiology, 4th ed. TheRoyal Society of Chemistry, Cambridge, United Kingdom.

4. Oyarzabal OA, Backert S. 2012. Microbial food safety. SpringerScience�Business Media, New York, NY.

5. Commission of the European Communities. 2005. Commission regula-tion (EC) 2073/2005 of 15 November 2005 on microbiological criteria forfoodstuffs. European Union, Brussels, Belgium. http://eur-lex.europa.eu/LexUriServ/LexUriServ.do?uri�OJ:L:2005:338:0001:0026:EN:PDF.

6. Li L, Xie B, Dong C, Wang M, Liu H. 2016. Can closed artificial ecosystemhave an impact on insect microbial community? A case study of yellowmealworm (Tenebrio molitor L.). Ecol Eng 86:183–189. https://doi.org/10.1016/j.ecoleng.2015.09.015.

7. EFSA Scientific Committee. 2015. Risk profile related to production andconsumption of insects as food and feed. EFSA J 13:4257. https://doi.org/10.2903/j.efsa.2015.4257.

8. Wynants E, Crauwels S, Verreth C, Gianotten N, Lievens B, Claes J, VanCampenhout L. 2018. Microbial dynamics during production of lessermealworms (Alphitobius diaperinus) for human consumption at industrialscale. Food Microbiol 70:181–191. https://doi.org/10.1016/j.fm.2017.09.012.

9. Varotto Boccazzi I, Ottoboni M, Martin E, Comandatore F, Vallone L,Spranghers T, Eeckhout M, Mereghetti V, Pinotti L, Epis S. 2017. A surveyof the mycobiota associated with larvae of the black soldier fly (Hermetiaillucens) reared for feed production. PLoS One 12:e0182533. https://doi.org/10.1371/journal.pone.0182533.

10. Zheng L, Crippen TL, Singh B, Tarone AM, Dowd S, Yu Z, Wood TK,Tomberlin JK. 2013. A survey of bacterial diversity from successive lifestages of black soldier fly (Diptera: Stratiomyidae) by using 16S rDNApyrosequencing. J Med Entomol 50:647– 658. https://doi.org/10.1603/ME12199.

11. Vandeweyer D, Crauwels S, Lievens B, Van Campenhout L. 2017. Met-agenetic analysis of the bacterial communities of edible insects fromdiverse production cycles at industrial rearing companies. Int J FoodMicrobiol 261:11–18. https://doi.org/10.1016/j.ijfoodmicro.2017.08.018.

12. Garofalo C, Osimani A, Milanovic V, Taccari M, Cardinali F, Aquilanti L,Riolo P, Ruschioni S, Isidoro N, Clementi F. 2017. The microbiota ofmarketed processed edible insects as revealed by high-throughputsequencing. Food Microbiol 62:15–22. https://doi.org/10.1016/j.fm.2016.09.012.

13. Dillon RJ, Dillon VM. 2004. The gut bacteria of insects: nonpathogenicinteractions. Annu Rev Entomol 49:71–92. https://doi.org/10.1146/annurev.ento.49.061802.123416.

14. Engel P, Moran NA. 2013. The gut microbiota of insects— diversity instructure and function. FEMS Microbiol Rev 37:699 –735. https://doi.org/10.1111/1574-6976.12025.

15. Jeon H, Park S, Choi J, Jeong G, Lee SB, Choi Y, Lee SJ. 2011. Theintestinal bacterial community in the food waste-reducing larvae ofHermetia illucens. Curr Microbiol 62:1390 –1399. https://doi.org/10.1007/s00284-011-9874-8.

16. Charlton AJ, Dickinson M, Wakefield ME, Fitches E, Kenis M, Han R,Zhu F, Kone N, Grant M, Devic E, Bruggeman G, Prior R, Smith R. 2015.Exploring the chemical safety of fly larvae as a source of protein foranimal feed. J Insects Food Feed 1:7–16. https://doi.org/10.3920/JIFF2014.0020.

17. Pinotti L, Ottoboni M, Giromini C, Dell’Orto V, Cheli F. 2016. Mycotoxincontamination in the EU feed supply chain: a focus on cereal byprod-ucts. Toxins (Basel) 8:45. https://doi.org/10.3390/toxins8020045.

Vandeweyer et al. Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 12

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from

18. Schneider JC. 2009. Principles and procedures for rearing high qualityinsects. Mississippi State University, Starkville, MS.

19. Vandeweyer D, Lenaerts S, Callens A, Van Campenhout L. 2017. Effect ofblanching followed by refrigerated storage or industrial microwavedrying on the microbial load of yellow mealworm larvae (Tenebriomolitor). Food Control 71:311–314. https://doi.org/10.1016/j.foodcont.2016.07.011.

20. Stoops J, Vandeweyer D, Crauwels S, Verreth C, Boeckx H, Van DerBorght M, Claes J, Lievens B, Van Campenhout L. 2017. Minced meat-likeproducts from mealworm larvae (Tenebrio molitor and Alphitobiusdiaperinus): microbial dynamics during production and storage. InnovFood Sci Emerg Technol 41:1–9. https://doi.org/10.1016/j.ifset.2017.02.001.

21. Klunder HC, Wolkers-Rooijackers J, Korpela JM, Nout MJR. 2012. Micro-biological aspects of processing and storage of edible insects. FoodControl 26:628 – 631. https://doi.org/10.1016/j.foodcont.2012.02.013.

22. Rumpold BA, Fröhling A, Reineke K, Knorr D, Boguslawski S, Ehlbeck J,Schlüter OK. 2014. Comparison of volumetric and surface decontamina-tion techniques for innovative processing of mealworm larvae (Tenebriomolitor). Innov Food Sci Emerg Technol 26:232–241. https://doi.org/10.1016/j.ifset.2014.09.002.

23. Fombong FT, Van Der Borght M, Vanden Broeck J. 2017. Influence offreeze-drying and oven-drying post blanching on the nutrient compo-sition of the edible insect Ruspolia differens. Insects 8:102–116. https://doi.org/10.3390/insects8030102.

24. Hanboonsong Y, Jamjanya T, Durst PB. 2013. Six-legged livestock: edibleinsect farming, collecting and marketing in Thailand. Food and Agricul-ture Organization of the United Nations, Regional Office for Asia and thePacific, Bangkok, Thailand.

25. Chao A. 1984. Non-parametric estimation of the number of classes in apopulation. Scand J Stat 11:265–270.

26. Shannon CE. 1948. A mathematical theory of communication. Bell SystTech J 27:379 – 423. https://doi.org/10.1002/j.1538-7305.1948.tb01338.x.

27. Krieg NR, Staley JT, Brown DR, Hedlund BP, Paster BJ, Ward NL, LudwigW, Whitman WB. 2010. Bergey’s manual of systematic bacteriology, vol4, 2nd ed. Springer Science � Business Media, New York, NY.

28. Brenner DJ, Krieg NR, Staley JT, Garrity GM, Boone DR, De Vos P,Goodfellow M, Rainey FA, Schleifer K-H. 2005. Bergey’s manual of sys-tematic bacteriology, vol 2, parts A, B and C, 2nd ed. Springer Science �Business Media, New York, NY.

29. Van Assche A, Álvarez-Pérez S, de Breij A, De Brabanter J, Willems KA,Dijkshoorn L, Lievens B. 2017. Phylogenetic signal in phenotypic traitsrelated to carbon source assimilation and chemical sensitivity in Acin-etobacter species. Appl Microbiol Biotechnol 101:367–379. https://doi.org/10.1007/s00253-016-7866-0.

30. Dahiru M, Enabulele OI. 2015. Incidence of Acinetobacter in fresh carrot(Daucus carota subsp. sativus). Int J Nutr Food Eng 9:1192–1195.

31. Hausdorf L, Fröhling A, Schlüter O, Klocke M. 2011. Analysis of thebacterial community within carrot wash water. Can J Microbiol 57:447– 452. https://doi.org/10.1139/w11-013.

32. Vandeweyer D, Crauwels S, Lievens B, Van Campenhout L. 2017. Micro-bial counts of mealworm larvae (Tenebrio molitor) and crickets (Achetadomesticus and Gryllodes sigillatus) from different rearing companies anddifferent production batches. Int J Food Microbiol 242:13–18. https://doi.org/10.1016/j.ijfoodmicro.2016.11.007.

33. Giaccone V. 2005. Hygiene and health features of “minilivestock,” p579 –598. In Paoletti MG (ed), Ecological implications of minilivestock:potential of insects, rodents, frogs and snails. Science Publishers, Inc.,Enfield, NH.

34. Grabowski NT, Klein G. 2016. Microbiology of processed edible insectproducts—results of a preliminary survey. Int J Food Microbiol 243:103–107. https://doi.org/10.1016/j.ijfoodmicro.2016.11.005.

35. Osimani A, Garofalo C, Milanovic V, Taccari M, Cardinali F, Aquilanti L,

Pasquini M, Mozzon M, Raffaelli N, Ruschioni S, Riolo P, Isidoro N,Clementi F. 2017. Insight into the proximate composition and microbialdiversity of edible insects marketed in the European Union. Eur Food ResTechnol 243:1157–1171. https://doi.org/10.1007/s00217-016-2828-4.

36. Banjo AD, Lawal OA, Adeyemi AI. 2006. The microbial fauna associatedwith the larvae of Oryctes monocerus. J Appl Sci Res 2:837– 843.

37. Ali A, Mohamadou BA, Siadou C, Aoudou Y, Tchiegang C. 2010. Physico-chemical properties and safety of grasshoppers, important contributorsto food security in the far north region of Cameroon. Res J Anim Sci4:108 –111. https://doi.org/10.3923/rjnasci.2010.108.111.

38. Zheng L, Crippen TL, Sheffield CL, Poole TL, Yu Z, Tomberlin JK. 2012.Evaluation of Salmonella movement through the gut of the lesser meal-worm, Alphitobius diaperinus (Coleoptera: Tenebrionidae). Vector BorneZoonotic Dis 12:287–292. https://doi.org/10.1089/vbz.2011.0613.

39. Templeton JM, De Jong AJ, Blackall PJ, Miflin JK. 2006. Survival of Campy-lobacter spp. in darkling beetles (Alphitobius diaperinus) and their larvae inAustralia. Appl Environ Microbiol 72:7909–7911. https://doi.org/10.1128/AEM.01471-06.

40. Magan N, Olsen M. 2004. Mycotoxins in food: detection and control, 1sted. CRC Press, Boca Raton, FL.

41. Jay JM, Loessner MJ, Golden DA. 2005. Modern food microbiology, 7thed. Springer Science � Business Media, New York, NY.

42. International Organization for Standardization. 2004. ISO 7932:2004. Micro-biology and food and animal feeding stuffs—horizontal method for theenumeration of presumptive Bacillus cereus—colony-count technique at30°C. International Organization for Standardization, Geneva, Switzerland.https://www.iso.org/standard/38219.html.

43. Stoops J, Crauwels S, Waud M, Claes J, Lievens B, Van Campenhout L.2016. Microbial community assessment of mealworm larvae (Tenebriomolitor) and grasshoppers (Locusta migratoria migratorioides) sold forhuman consumption. Food Microbiol 53:122–127. https://doi.org/10.1016/j.fm.2015.09.010.

44. Dijk R, van den Berg D, Beumer R, de Boer E, Dijkstra A, Mout L,Stegeman H, Uyttendaele M, in ’t Veld S. 2015. Microbiologie vanvoedingsmiddelen: methoden, principes en criteria, 5th ed. MYbusiness-media, Capelle aan den Ijssel, The Netherlands.

45. Lievens B, Brouwer M, Vanachter ACRC, Lévesque CA, Cammue BPA,Thomma BPHJ. 2003. Design and development of a DNA array for rapiddetection and identification of multiple tomato vascular wilt pathogens.FEMS Microbiol Lett 223:113–122. https://doi.org/10.1016/S0378-1097(03)00352-5.

46. Benson DA, Cavanaugh M, Clark K, Karsch-Mizrachi I, Lipman DJ, OstellJ, Sayers EW. 2013. GenBank. Nucleic Acids Res 41:D36 –D42. https://doi.org/10.1093/nar/gks1195.

47. International Organization for Standardization. 2017. ISO 6579-1:2017. Mi-crobiology of the food chain—horizontal method for the detection, enu-meration and serotyping of Salmonella—part I: detection of Salmonella spp.International Organization for Standardization, Geneva, Switzerland. https://www.iso.org/standard/56712.html.

48. Association Française de Normalisation. 1998. AFNOR BRD 07/4-09/98. RAPID’L.mono (Detection). Association Française de Normalisa-tion, La Plain Saint-Denis, France. https://nf-validation.afnor.org/wp-content/uploads/sites/2/2014/03/BRD-07-04-09-98_en.pdf.

49. Association Française de Normalisation. 2003. AFNOR 3M 01/9-04/03 B.3M Petrifilm Staph Express Count System. Association Française deNormalisation, La Plain Saint-Denis, France. https://nf-validation.afnor.org//en/wp-content/uploads/sites/2/2014/03/3M-01-09-04-03-B_en.pdf.

50. Edgar RC. 2016. SINTAX: a simple non-Bayesian taxonomy classifier for16S and ITS sequences. bioRxiv. https://doi.org/10.1101/074161.

51. R Development Core Team. 2013. R: a language and environment forstatistical computing. R Foundation for Statistical Computing, Vienna,Austria.

Microbial Dynamics of a Cricket-Rearing Cycle Applied and Environmental Microbiology

June 2018 Volume 84 Issue 12 e00255-18 aem.asm.org 13

on October 23, 2020 by guest

http://aem.asm

.org/D

ownloaded from