# Amylase activity across black soldier fly larvae development and feeding substrates: insights on starch digestibility and external digestion

J.B. Guillaume, J.L. da Lage, S. Mezdour, Frédéric Marion-Poll, C M C

Terrol, C.M.C. Brouzes, Philippe Schmidely

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J.B. Guillaume, J.L. da Lage, S. Mezdour, Frédéric Marion-Poll, C M C Terrol, et al.. Amylase activity across black soldier fly larvae development and feeding substrates: insights on starch digestibility and external digestion. Animal, 2024, 18 (11), pp.101337. 10.1016/j.animal.2024.101337. hal-04706044

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Amylase activity across black soldier fly larvae development and feeding substrates: insights on starch digestibility and external digestion

J.B. Guillaume, J.L. Da Lage, S. Mezdour, F. Marion-Poll, C. Terrol, C.M.C. Brouzes, P. Schmidely

PII: S1751-7311(24)00274-X

DOI: https://doi.org/10.1016/j.animal.2024.101337

Reference: ANIMAL 101337

To appear in: *Animal*

Received Date: 15 March 2024

Revised Date: 7 September 2024

Accepted Date: 13 September 2024

Please cite this article as: J.B. Guillaume, J.L. Da Lage, S. Mezdour, F. Marion-Poll, C. Terrol, C.M.C. Brouzes, P. Schmidely, Amylase activity across black soldier fly larvae development and feeding substrates: insights on starch digestibility and external digestion, *Animal* (2024), doi: https://doi.org/10.1016/j.animal.2024.101337

This is a PDF file of an article that has undergone enhancements after acceptance, such as the addition of a cover page and metadata, and formatting for readability, but it is not yet the definitive version of record. This version will undergo additional copyediting, typesetting and review before it is published in its final form, but we are providing this version to give early visibility of the article. Please note that, during the production process, errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

© 2024 Published by Elsevier B.V. on behalf of The Animal Consortium.

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**Full title**

Amylase activity across black soldier fly larvae development and feeding substrates: insights on starch digestibility and external digestion

**Authors**

J.B. Guillaume<sup>1,2,3*</sup>, J.L. Da Lage<sup>2</sup>, S. Mezdour<sup>4</sup>, F. Marion-Poll<sup>2,5</sup>, C. Terrol<sup>3</sup>, C.M.C. Brouzes<sup>3</sup> and P. Schmidely<sup>1,5</sup>

<sup>1</sup> Université Paris-Saclay, INRAE, AgroParisTech, UMR Modélisation Systémique Appliquée aux Ruminants, 91120 Palaiseau, France

<sup>2</sup> Laboratoire Évolution, Génomes, Comportement et Écologie, CNRS, IRD, Université Paris-Saclay, Institut Diversité, Ecologie et Evolution du Vivant (IDEEV), 91190 Gif-sur-Yvette, France

<sup>3</sup> Agronutris, R&D Department, 31650 Saint-Orens de Gameville, France

<sup>4</sup> Université Paris-Saclay, UMR Sayfood, AgroParisTech, INRAE, 91120 Palaiseau, France

<sup>5</sup> Université Paris-Saclay, AgroParisTech, 91120 Palaiseau, France

**Corresponding author:**

Jérémy B. Guillaume. E-mail: jeremy.guillaume@agronutris.com

**Highlights**

* Mechanisms of starch digestion in black soldier fly larvae are poorly understood

* Larval amylase activity increased over time and dropped when approaching pupation

* Substrate starch and sugars contents did not affect larval amylase activity

* Reducing sugars absorbed by the larvae could repress amylase activity

* Black soldier fly larvae could have an external digestive activity
**Abstract**

Black soldier fly larvae (BSFL ; *Hermetia illucens*) hold promise for converting biowaste into proteins and lipids for feed. Dietary starch is efficiently digested by the larvae and influences larval performance, but the mechanisms of starch digestion remain poorly understood. This study investigated changes of individual weight and amylase activity in BSFL after 4, 7 and 11 days of feeding for five substrates varying in starch content and type: chicken feed (CF), corn gluten feed (CGF), wheat bran (WB), wheat distillers grain (WDG) and discarded potatoes (DP). Substrate amylase activities were also measured with and without larvae (feeding and fermenting trays, respectively) over time in order to explore external digestion. Feed conversion ratio (FCR) and estimated digestibility (ED) of dry matter and starch were assessed at the end of the experiment. The ranking for best FCR was: CF, WB, CGF, WDG and DP. In feeding trays, ED of dry matter was 69.8 ± 1.8%, 59.5 ± 2.9%, 58.6 ± 0.7%, 45.4 ± 0.6% and 19.5 ± 0.8% in CF, DP, WB, CGF and WDG, respectively. Estimated digestibility of starch reached 100% with WB and CGF, followed by CF (88.2 ± 2.3%), DP (85.2 ± 1.2%) and WDG (43.1 ± 1.0%). Larval amylase activity increased with growth for all substrates, and dropped when approaching pupation. No relationship was found between larval amylase activity and substrate starch or other nutrients content, but a negative correlation was reported with the reducing sugars content of the larvae, suggesting glucose repression of amylase production. Amylase activity decreased with time in all feeding and fermenting substrates except WDG and DP. *In vitro* degradation assays indicated that BSFL amylase was nine times more efficient on raw corn or wheat starch than on raw potato starch, highlighting that starch structure is a major driver of digestibility. Western blot analysis revealed the presence of BSFL amylase in the feeding substrate, hinting at external digestion. Larval amylase was purified to identify its optimal pH (5.0-6.5) and temperature (70°C). These results highlight that starch content is not a major driver of amylase activity in BSFL and suggest that other non-investigated factors could have had a crucial impact on the activity of larval digestive enzymes, such as microbial community of the substrate and presence of amylase inhibitors. This study also provides insights into the evolution of BSFL digestive activity during their development and the occurrence of external digestion.

**Keywords**

*Hermetia illucens* ; digestive enzymes ; bioconversion ; insects as feed ; Diptera

**Implications**

Substrate starch content is a major driver of black soldier fly larvae performance, but the mechanisms of starch digestion are poorly understood. This study investigated larval and substrate amylase activity over time in five substrates varying in starch content. Results showed that larval amylase activity increased during growth and
dropped in preparation for pupation. No correlation between larval amylase activity and starch or other nutrient contents of the substrate was identified. Amylase activity decreased over time in most substrates, but amylase from black soldier fly larvae was detected in consumed substrates, suggesting that the larvae have an external digestion.

Introduction

In the last decades, the black soldier fly (BSF; *Hermetia illucens* (L.); Diptera: Stratiomyidae) has been extensively studied for the ability of its larvae (**BSFL**) to feed on an extremely large range of organic substrates including agricultural by-products (Broeckx et al., 2021), catering waste (Cho et al., 2020), slaughterhouse waste (Naser El Deen et al., 2023) and human or animal faeces (Diener et al., 2011; Rehman et al., 2019). These larvae are rich in proteins, lipids (Barragan-Fonseca et al., 2017) and antimicrobial compounds (Gasco et al., 2021) and therefore show great potential for use as feed for poultry (Martínez Marín et al., 2023), pig (Veldkamp and Vernooij, 2021), fish (Weththasinghe et al., 2021), or pets (Kotob et al., 2022). Current major sources of proteins for animal feed (i.e. soybean and fish meal) are severely criticised for their negative impacts on the environment (Hall, 2010; Jia et al., 2020), and BSFL production holds promises both for improving the sustainability of feed protein supply (Smetana et al., 2019) and treating biowaste (Tchonkouang et al., 2023). However, BSFL feed conversion efficiency and nutritional content greatly depend on the substrate they are fed (Gold et al., 2018; Surendra et al., 2020; Hopkins et al., 2021). Recent studies identified that digestible carbohydrate (mostly starch and sugars) content was one of the main drivers of larval performance (Barragan-Fonseca et al., 2021; Cheon et al., 2021) and that BSFL were highly efficient in digesting starch (Guillaume et al., 2023).

Whether starch digestion may limit substrate digestibility and larval growth remains undetermined. Starch is a polysaccharide found in plants constituted by glucose units linked by α-1,4-glycosidic bonds in its linear parts and α-1,6-glycosidic bonds at the branches. Amylose and amylopectin are the linear and branched fractions of starch, respectively (Junejo et al., 2022). Glycosidic linkages in starch can be broken down by alpha- (EC 3.2.1.1; in animals, plants, bacteria and fungi), beta- (EC 3.2.1.2; in plants, bacteria and fungi) and gamma-amylases (EC 3.2.1.3; in plants and animals) thus releasing reducing sugars (Bairoch, 2000; Akinfemiwa et al., 2023). A query on NCBI returned that the genome of *H. illucens* (published by Generalovic et al., 2021, accession number GCF_905115235.1) had nine genes coding for alpha-amylase-like
proteins. Most Diptera species only have two to six (Da Lage, 2018), highlighting the importance of starch digestion in black soldier fly metabolism. Microorganisms from the substrate and the larval gut also produce alpha-amylase and participate in starch digestion (Muharram et al., 2021; Bui Thi Cam et al., 2023). Unless specified, in the present paper, the term ‘amylase’ will be used to refer indifferently to enzymes from BSFL (alpha-amylase), plant material of the feeding substrate (alpha-, beta- or gamma-amylase) or microbes of the larval gut or the substrate (alpha- or beta-amylase). Previous research identified that starch digestion mainly occurs in the anterior midgut of BSFL (Bonelli et al., 2019) and that this organ could adapt its morphological and transcriptional features depending on the diet ingested by the larva (Bonelli et al., 2020). Gut amylase activity varies with the feeding substrate (Sun et al., 2022; Chen et al., 2023), and although diet carbohydrate content is hypothesised to be a major driver of amylase production (Bonelli et al., 2020), it remains to be established if amylase activity is correlated with starch or sugar content of the substrate. Many other aspects of starch digestion in BSFL remain to be explained (Tettamanti et al., 2022), including the evolution of amylase activity over larval development. Virtually all the studies exploring BSFL amylase activity used larvae of late development stages, i.e. 19-day-old individuals (Sun et al., 2022), 5th instar larvae (Kim et al., 2011) or 6th instar larvae (prepupae) (Kim et al., 2010; Bonelli et al., 2020; Chen et al., 2023). To the best of our knowledge, only Intayung et al., 2021 measured amylase activity over time (6, 12 and 18-day-old larvae), for BSFL fed a single substrate. The interaction between substrate nutritional content and larval development time on BSFL amylase activity remains to be explored.

Black soldier fly larvae are known to aggregate in clusters (Shishkov et al., 2019; Wu et al., 2023), producing heat due to crawling (Li et al., 2023), which improves their collective feed efficiency at high density (Guillaume et al., 2023). This behaviour is called ‘social digestion’ and is often associated with ‘external digestion’ in insects (release of digestive enzymes in the environment before ingestion), which has been suspected in BSFL (Gold et al., 2018) but never demonstrated. External digestion has been documented in various saprophagous Diptera larvae such as *Drosophila melanogaster* (Haj-Ahmad and Hickey, 1982; Gregg et al., 1990; Sakaguchi and Suzuki, 2013), *Lucilia sericata* (Hobson, 1931) and *Lucilia cuprina* (Mackerras and Freney, 1933). Recently, external digestion was described in *Anastrepha ludens* larvae by measuring an increase of enzymatic activity in the feeding substrate, both over time and larval density (Aceituno-Medina et al., 2023). This approach is promising to identify whether BSFL have an external digestion activity.

The present study addresses the question of post-ingestion regulation of amylase activity in BSFL depending on starch content, sugar content and starch type of the feeding substrate, and relates it with larval growth and starch digestion. Purified BSFL amylase is produced to determine the optimal pH and temperature of this enzyme. Finally, our work uses two different approaches (substrate amylase activity and Western blot) to explore the role of amylases in the substrate and shed light on external digestion in BSFL.
# Material and Methods

## Animals

The Agronutris R&D facility (Toulouse, France) supplied BSFL. The strain has been reared for over 50 generations on an industrial feeding substrate composed of agricultural by-products mostly from the grain and tuber industries. For experimental purposes, 400g of chicken feed mash (CF) (Maïsadour, Sud-Ouest Aliment VT100 Poulet Jaune Démarrage, Haut-Mauco, France) mixed with warm water to reach a 64 ± 0.5% moisture content were placed in a 12×10×7 cm plastic container, topped with 5 to 10 g of wheat bran to absorb excess moisture, and inoculated with 0.6 g of neonate larvae. Three identical nursery boxes were prepared and closed with a mosquito net and a cut lid. They were maintained at 28°C and 65% relative humidity for five days, before being shipped to the experimental laboratory in less than 24 hours. The content of all boxes was then mixed in a 32×20 cm container and kept at 25°C for one more day to obtain 7-day-old starter larvae. To standardise their size and remove remaining substrate (frass) from the nursery, starter larvae were successively sieved using screens with 2.4 and 1.0 mm mesh size. Three samples of approximately 1 g of larvae were weighed and counted manually to calculate the average fresh weight of a starter larva (3.7 ± 0.1 mg), that is later used to calculate the weight of starter larvae that should be inoculated.

## Feeding substrates and nutritional parameters analysis

Five feeding substrates varying in starch, carbohydrate (defined as the sum of starch and simple sugars contents), crude protein and protein+carbohydrate contents, and protein:carbohydrate ratio were used: CF (the same that was used during the nursery period), corn gluten feed (CGF), wheat bran (WB), wheat distillers grain (WDG) and discarded potatoes (DP). Chicken feed was used as a standard diet while the four others were agricultural by-products potentially relevant for the formulation of a compound BSFL diet on an industrial scale and produced in the North-East of France (purchased from Néalia, Pauvres, France). The substrates were also selected to cover different starch vegetal origins: starch comes from various plants in CF (wheat, corn, soybean, rapeseed), corn in CGF, wheat in WB and WDG, and potato in DP. Dry matter analyses with a thermobalance set at 120°C (Radwag MA 50.R, Radom, Poland) indicated that CF, WDG and WB had low initial moisture content (7.6%, 9.0% and 7.8% respectively) and could be stored at room temperature with no risk of decaying, while CGF and DP contained more water (59.7% and 74.0% respectively) and were kept frozen at -20°C between reception and use.

Each substrate was mixed with warm tap water using a hand blender (Electrolux Professional Bermixer Pro 450 W, Saint-Denis, France; maximum power for 2:30 min). Targeted moisture contents were 68% for CF and WDG, 78% for CGF and WB, and 74% for DP. These values had been validated during preliminary trials to obtain similar textures suitable for BSFL ingestion and growth. As DP did not require water addition, it was kept in a 25°C climate-controlled room for 24 h before use to avoid difference in the initial temperature of the substrates, and then mixed with the hand blender.
Substrates were sampled right before use. They were oven-dried at 60°C until constant weight and sent for analysis to Upscience (Saint-Nolff, France): nutritional content was assessed enzymatically for starch (internal method inspired by NF V18-121 - March 1997 repealed), using the Luff-Schoorl technique for total sugars (expressed as glucose; internal method inspired by CE 152/2009) and the Dumas method (internal method inspired by NF EN ISO 16634-1) with a conversion factor of 6.25 for crude protein. Table 1 presents the nutritional parameters of each substrate on a DM basis.

Experimental conditions and sampling of larvae and substrates

Assays were performed in 16.5×10.6×7 cm plastic trays (Pro-Jet BG110, Saint-Ouen-l'Aumône, France). For each substrate, five feeding trays were prepared using 420 g of fresh substrate inoculated with 500 starter larvae. Five fermentation trays were also started with the same weight of substrate but without larvae, to compare substrate amylase activity and starch degradation with and without larvae. Feeding and fermentation trays will respectively be referred to as ‘CF+’ and ‘CF-’, for the example of CF. All trays were placed in a light pantyhose and closed with a lid with a 16×10 cm opening to avoid the escape of larvae while enabling respiration. Feeding and fermentation trays were maintained at 28 ± 0.5°C, 75 ± 2% relative humidity, and with a L12:D12 photoperiod, in the same climatic chamber (Percival Scientific DR-36VL, Perry, GA, USA). Trays were randomly relocated daily to avoid possible effects of local climatic conditions.

After 4, 7 and 11 days of experiment (D04, D07 and D11, respectively), one tray of each feeding and fermenting substrate was randomly selected for sampling (Fig. 1). In feeding trays, larvae were removed from the substrate with forceps, rinsed with lukewarm tap water, and dried with a paper towel. Thirty larvae were individually weighed on a precision scale (± 0.1 mg) to calculate their average fresh weight. For each substrate, three samples of 10 individuals (or more in order to have at least 1.0 g of larvae) were collected and frozen at -80°C for further homogenisation and amylase activity analysis. Substrates were thoroughly inspected for the absence of larvae and three samples of 1.0 g were collected and frozen. Four grams of larvae and substrate were used to measure their respective DM content with a thermobalance as previously described. As fermentation trays contained no larvae, only substrate was sampled and analysed. Trays used for sampling on day 4 and 7 were terminated. On D11, the three remaining trays of each treatment were weighed and all their larvae were meticulously removed, rinsed, dried and weighed separately before being killed by freezing at -20°C. After sampling, the substrate of each tray was entirely collected, oven-dried at 60°C until constant weight and analysed for starch content separately with the same technique as initial substrates. Feed conversion ratio (FCR) was calculated for each D11 tray that had received larvae as the ratio between the initial dry weight of substrate and total dry larval weight gain. To account for substrate loss due to larvae rinsing, final substrate weight was defined as final tray weight, minus final weight of the larvae and weight of empty tray. The FCR accounts for the efficiency of conversion of the substrate into total larval biomass, with a lower FCR indicating a more efficient bioconversion. Estimated digestibility (ED) of DM was calculated for
each D11 tray as the difference between initial and final substrate weight divided by initial substrate weight (on a dry basis) according to Guillaume et al., 2023. Initial and final starch contents were used to calculate ED of starch correspondingly. For samples where starch could not be detected (detection threshold: 0.6% DM), we considered that no starch remained in the substrate, hence the associated ED of starch was 100%.

## Homogenates preparation

For each condition of substrate and experimental time, three homogenates (technical replicates) of larvae, feeding substrates and fermenting substrates each were prepared. Frozen samples of larvae and substrates were thawed at 4°C and then placed in 3.0 mL cold phosphate-buffered saline. Thirty microliters of protease inhibitor (Halt Protease Inhibitor Cocktail, Thermo Scientific 78429, Waltham, MA, USA) were added to prevent amylase degradation by digestive protease from the sample. To avoid sample melanisation due to the release of phenoloxidase from the larval hemolymph, a few crystals of N-phenylthiourea (Sigma-Aldrich P7629, St. Louis, MO, USA), a phenoloxidase inhibitor (Ryazanova et al., 2012), were added. Samples were ground at 1°C in 7 mL tubes with 2.8 mm ceramic beads (Bertin Technologies P000935-LYSK0-A, Montigny-le-Bretonneux, France) using a bead mill coupled to a dry ice cooling system (Precellys Evolution Touch and Cryolys, Bertin Technologies; 30 s, 8 500 rpm). Preparations were then centrifuged for 5 min at 29 500 g, 4°C, floating particles and pellet were eliminated and a second centrifugation was performed to collect a homogeneous liquid phase. Homogenates were kept at -20°C until further analysis.

## Amylase optimal pH and temperature determination

One homogenate of larvae fed CF for 7 days was further purified. All the following steps were performed at low temperature (cold room at 4°C, sample preserved in ice between different steps) on a magnetic stirrer. Solid ammonium sulfate was added to the homogenate up to 80% saturation (2.40 g for 4.5 mL of homogenate) in order to neutralise protein charges and precipitate them. The solution was stirred for two hours and then centrifuged at 12 000 g and 4°C for 50 min. The protein pellet was collected and manually solubilised in HEPES buffer (20 mmol/L HEPES, 20 mmol/L NaCl, 1 mmol/L CaCl2, pH=7.5). Considering that Diptera amylases have a molecular weight of about 50 kDa (Da Lage, 2018), proteins and other molecules of low molecular weight were then eliminated by performing an extensive dialysis (18 hours) against 450 mL HEPES buffer using a membrane with a 12-14 kDa cutoff (Spectra/Por Dialysis Membrane 4, Repligen 132706, Waltham, MA, USA). Amylases were further purified according to Schramm and Loyter, 1966: nucleic acids were precipitated by slowly adding pure -20°C ethanol to bring the solution to a 40% ethanol concentration, stirring for 40 min, and then centrifuging at 48 000 g, 4°C for 40 min. About 7 mL of supernatant were then collected and used for selective amylase precipitation with 400 µL of a 16 mg/mL glycogen solution (Sigma-Aldrich G8751) with agitation for 20 min. After centrifuging 20 min at 48 000 g, 4°C, the pellet composed of an amylase-
glycogen complex was collected, dissolved in 250 µL HEPES, and left for two hours at room temperature to allow glycogen degradation by amylases. Finally, a second dialysis was performed in the same conditions as the first one to eliminate sugars released from glycogen hydrolysis. The resulting purified amylase solution was then used for optimal pH and temperature determination.

Amylase activity was assessed according to Bernfeld, 1955: potato starch (Sigma-Aldrich 33615) was used as a substrate and reducing sugar production was revealed by measuring 540 nm absorbance increase with 3,5-dinitrosalicylic acid (DNS). To identify optimal pH, twelve buffer solutions (20 mmol/L NaCl and 1 mmol/L CaCl2) with pH ranging from 3.0 to 8.5 and starch concentration of 10 mg/mL were prepared. Starch was solubilised by bringing each solution to a boil with constant stirring. Two hundred microliters of starch solution were added with 20 µL of purified amylase solution in a 1.5 mL tube, vortexed, and put in a 37°C dry bath incubator for exactly 5:00 min for starch hydrolysis. Two hundred microliters of DNS were then added, and the tube was vortexed and put at 100°C. After 5:00 min, the tube was placed in an ice bath for 5:00 min. Finally, 1.0 mL of distilled water was added and mixed by pipetting, and absorbance was immediately measured against a blank with distilled water instead of amylase solution. Six technical replicates were assayed per condition for pH values between 6.0 and 7.0, and three technical replicates for other pH values. A similar approach was used to determine optimal temperature: we used 10 mg/mL potato starch solution at pH 6.0 and seventeen different reaction temperatures ranging from 15 to 95°C, with a 5:00 min incubation time. Six technical replicates were used at 65, 70 and 75°C, and three technical replicates for other temperatures.

## Larvae and substrates amylase activity determination

First, the soluble protein concentration of each homogenate was determined in triplicate by measuring its absorbance at 595 nm in the presence of Coomassie blue (Protein Assay Dye Reagent Concentrate, Bio-Rad 5000006, Hercules, CA, USA) and using BSA as standard according to Bradford, 1976. Amylase activity was measured in duplicate spectrophotometrically at 540 nm with DNS as described above, using 10 mg/mL potato starch in pH 6.0 MES buffer (20 mmol/L MES, 20 mmol/L NaCl and 1 mmol/L CaCl2) and an incubation temperature of 37°C. However, as larvae and substrate homogenates could contain reducing sugars, absorbance was also assayed in duplicate with 200 µL MES buffer, 20 µL sample, and 200 µL DNS (sample blank), and the mean result was subtracted from absorbance with starch. We also subtracted the absorbance due to reducing ends of the native starch solution (starch solution blank) (Supplementary Figure S1). For some substrate samples with very low amylase activity, this calculation led to negative results: in that case, we considered that no amylase activity was detected. Amylase activity was expressed as the number of micromoles of maltose released per second and per gram of soluble protein in the homogenate (hereafter noted U). Three homogenates were assayed for each experimental treatment.
**In vitro enzymatic efficiency assays on different types of raw starch**

To gain insight into the enzymatic efficiency of BSFL amylase on different types of starch, we measured BSFL amylase activity on raw starch (i.e. unmodified commercially available starch) of the same plant origin as the different feeding substrates. Raw starch from rice was also included in our study as BSFL can be reared on wasted rice (Ong et al., 2017), rice hulls (Guo et al., 2021) or rice straw (Manurung et al., 2016; Liu et al., 2021). Suspensions of 10 mg/mL raw starch from corn, wheat, potato and rice (Sigma-Aldrich S9679, S5127, S4251 and S7260, respectively) were prepared in pH 6.0 MES buffer (20 mmol/L MES, 20 mmol/L NaCl and 1 mmol/L CaCl₂) by stirring at room temperature. Amylase activity of a BSFL homogenate (larvae fed CF for 7 days) on each type of raw starch was measured using the Bernfeld technique. The hydrolysis step was performed with 250 µL of raw starch suspension and 25 µL of homogenate in a dry bath incubator at 37°C with continuous stirring (900 rpm) to maintain starch granules in suspension. After 5 hours, tubes were placed in an ice bath for a few minutes to stop the reaction and centrifuged for 4 min at 14 000 rpm to pellet the remaining starch. Two hundred and twenty microliters of supernatant were pipetted in new tubes and treated with 200 µL of DNS at 100°C for exactly 5 min. Tubes were then placed in an ice bath for exactly 5:00 min and 540 nm absorbance was measured against a blank of 250 µL MES buffer added with 25 µL distilled water and treated in the same conditions. We used four technical replicate tubes per type of raw starch. Reducing sugars from the larvae homogenate and spontaneous hydrolysis of raw starches were assayed in triplicate likewise and subtracted from the results. Enzymatic efficiency results were expressed in U as well.

**Microscopic observation of starch granules**

Initial substrates and raw starches were mounted between a slide and a coverslip in distilled water and added with 10% Lugol for initial substrates. Slides were observed with a microscope coupled with a camera (Leica DM6000 B and DFC490, Wetzlar, Germany) to qualitatively describe the size and shape of starch granules.

**Detection of black soldier fly larvae amylase by Western blot**

The presence of amylase from BSFL in consumed substrate was investigated by Western blot. We used homogenates from larvae and substrates obtained with CF, and a *D. melanogaster* amylase purified at the lab as a positive control. Proteins from each homogenate were denatured using LDS (NuPAGE LDS Sample Buffer, Invitrogen NP0007, Waltham, MA, USA), heated at 95°C for 3 min, and then loaded on a 4-12% Bis-Tris gel (Invitrogen NP0322BOX). Each well received a standardised amount of 10 µg protein using concentrations determined previously with the Bradford technique, except for the purified *D. melanogaster* amylase load that was only 75 ng. SeeBlue Plus2 Prestained Standard (Invitrogen LC5925) was used as the molecular weight marker in the first well. Proteins were separated by electrophoresis with MOPS SDS as a running buffer (Invitrogen NP0001) and then electro-transferred on a nitrocellulose membrane (iBlot Gel Transfer Stacks Nitrocellulose Mini and iBlot Dry
Blotting System, Invitrogen IB301002 and IB1001). The membrane was then successively incubated at room temperature on an agitating plate with 30 mL PBS for 10 min to rinse it, with 30 mL of 5% defatted powdered milk in PBS-Tween for 90 min to block the membrane, and with 10 mL of 1/2 500 primary antibody and 1% defatted milk in PBS-Tween for one hour at room temperature before being stored overnight at 4°C. The primary antibody, raised against *Drosophila* amylase, had been produced in guinea pig by Eurogentec (Liege, Belgium) from purified recombinant *D. melanogaster* amylase produced in the laboratory for previous research. The membrane was then rinsed four times with 30 mL of PBS-Tween for 15 min, incubated with 1/5 000 secondary antibody (anti-guinea-pig produced in goat, Sigma-Aldrich A7289) and 1% defatted milk in PBS-Tween for 60 min, rinsed three times with 30 mL of PBS-Tween for 20 min and one last time with 30 mL of PBS for 15 min. The membrane was finally revealed with a chemiluminescent detection kit (Cytiva RPN2232, Washington D.C., USA) and immediately recorded with an imager (ChemiDoc Imaging System, Bio-Rad). Lanes and bands were automatically detected on an uncropped unprocessed image of the membrane using Image Lab (Image Lab 6.1.0, Bio-Rad, 2020). Two Western blots were assessed on different homogenates from CF larvae and substrates.

## Statistical analysis

Data were analysed using the R software (version 4.3.2, R Core Team, 2023). Plots and figures in the text represent the mean ± standard error of the mean. To assess the effect of categorical factors on investigated quantitative variables, GLM were used with a gaussian distribution when residuals were normally distributed according to a Shapiro-Wilk test, or with a gamma distribution. Post-hoc pairwise comparisons were performed with a Bonferroni adjustment, except for the comparisons between maximal relative amylase activity and activity obtained in other pH or temperature conditions, that were assessed with a Dunnett test. The significance level used for the tests was 5%. The number of replicates is indicated below each figure. We explored the influence of initial substrate nutritional parameters (starch, sugar, carbohydrate, crude protein and protein+carbohydrate contents, protein:carbohydrate ratio) on the amylase activity of larvae, feeding substrates and fermenting substrates (overall and for each sampling day) with linear regressions. We also assayed potential correlations between homogenates reducing sugar content (measured with the Bernfeld approach using MES buffer without starch added with homogenate, as presented in Supplementary Figure S1) and amylase activity, as well as between amylase activity in larvae and in substrate from feeding trays, and between amylase activity in substrates from feeding trays and fermenting trays. The code for the statistical analysis is available as a Supplementary Material S1.
# Results

## Larval growth

Feeding substrate, time and their interaction had an effect on individual larval fresh weight. All feeding substrates were associated with larval growth (Fig. 2). Maximal individual weight was reached with CF on D11 with 203.7 ± 6.1 mg. However, no weight gain was observed with this substrate between D07 and D11. Similarly, BSFL fed CGF or WB had no weight variation between D07 and D11. This is associated with the start of the non-feeding stage, as dark prepupae were observed on D11 in these three substrates. Contrarily, no prepupae were present in WDG or DP and larvae gained weight continuously from the start of the experiment to D11 when fed these substrates, but reached lower maximal weight (85.9 ± 3.5 mg and 82.1 ± 6.1 mg, respectively). Larval DM content after 11 days of feeding varied between substrates (Supplementary Figure S2). The best-performing substrate was CF with an FCR of 3.92 ± 0.02, followed by WB, CGF, WDG and DP (Fig. 3). These results are consistent with individual weight on D11.

## Estimated digestibility of dry matter and starch

Substrate, presence of larvae and their interaction had an effect on ED of DM and ED of starch (Figs. 4.A and 4.B). In all substrates except CGF, both DM and starch were more digested with larvae than without. Maximal ED of DM with larvae was reached in CF (69.8 ± 1.8%). In fermenting trays, CGF and DP had the highest ED of DM (38.8 ± 0.4% and 38.5 ± 1.3%, respectively). For ED of starch with larvae, CGF and WB ranked first with a noticeable value of 100% in the three trays of these two substrates. Maximal ED of starch without larvae was also reached in CGF (94.4 ± 3.3%) but was significantly lower in WB (26.8 ± 1.9%). Both with and without larvae, WDG had the lowest ED of DM (19.5 ± 0.8% and 3.6 ± 0.7%, respectively) and ED of starch (43.1 ± 1.0% and 6.5 ± 3.2%, respectively).

## Larval amylase optimal pH and temperature

Maximal amylase activity was observed at pH 6.5 (Fig. 5.A), but it plateaued above 90% of the maximal activity for pH varying between 5.0 and 6.5. Amylase activity was negligible in both extreme pH conditions investigated (3.0 and 8.5). Results of temperature assays revealed a large thermal range of enzymatic activity, as relative amylase activity was above 50% between 40 and 75°C (Fig. 5.B). Amylase activity peaked at 70°C and was null both at 15 and 95°C. It was decided to investigate amylase activity of larvae and substrates homogenates at pH 6.0 to concur with the pH of the anterior midgut (major place of amylase production in BSFL) reported by Bonelli et al., 2019, and at 37°C as the optimal temperature of 70°C was not biologically relevant.
**Larval amylase activity**

Larval amylase activity varied with feeding substrate, sampling day and their interaction. Maximal amylase activity in BSFL was reached on D07 in WB (29.3 ± 3.4 U), CF (23.9 ± 2.1 U) and CGF (16.9 ± 0.4 U), and on D11 with DP (18.9 ± 2.8 U) and WDG (16.1 ± 2.4 U) (Fig. 6). Amylase activity in starter larvae was 15.7 ± 0.2 U. No correlation was found between amylase activity in larvae homogenates and the nutritional parameters of initial substrates, both considering all sampling days or each sampling day separately (Supplementary Table S1). Larval amylase activity and reducing sugar content of the associated substrate homogenates were also not correlated. However, we found a negative correlation between amylase activity in larvae and the reducing sugar content of these homogenates (Fig. 7).

**Substrate amylase activity**

Amylase activity of substrate was affected by sampling day in CF, CGF and WB but not in WDG and DP, and by presence of larvae in CF and WB only (Fig. 8). The interaction of both parameters had an effect in CF, WB and CGF. Initial substrates (D00) either displayed the highest amylase activity (CF, WB and WDG) or a value non-significantly different from the highest (CGF, DP). The variation range of substrate amylase activity greatly differed between substrates. Both for consumed and fermented substrate samples, no correlation was found between homogenates amylase activity and initial substrates nutritional parameters (considering all samples together, or samples from each day separately), or homogenates reducing sugar content. Amylase activity of substrate homogenates from feeding trays was not correlated with amylase activity of associated larvae homogenates, nor with amylase activity of associated substrates from fermented trays (Supplementary Table S1).

**Enzymatic efficiency of black soldier fly larvae amylase on different types of raw starch**

No difference was detected between amylase efficiency on corn and wheat starch, but they were 9 times more efficiently hydrolysed than potato starch (Fig. 9). Amylase activity on rice starch was 1.5 times higher than on corn or wheat starch.

**Microscopic observation of starch granules**

Starch granules were bigger in potato (up to 150 µm in DP and 50 µm in raw potato starch) than in wheat (5-30 µm) or corn (10-20 µm), while the smallest were in rice (less than 5 µm) (Supplementary Figures S3 and S4). These observations, along with shape differences, are summarised in Table 2.
Detection of black soldier fly larvae amylase by Western blot

Amylase was detected in larvae homogenates from D04, D07 and D11 (Fig. 10), with bands varying in intensity accordingly with amylase activity detected by the Bernfeld technique (Fig. 6). Amylase in BSFL had the same molecular weight as the control amylase from *D. melanogaster*, evaluated around 44 kDa. Faint bands around 41 kDa were also detected in D04 and D07 larvae, and around 59 kDa in D07 and D11 larvae. No signal could be detected in initial and fermented substrates. This confirms that the used antibody is binding specifically to insect or Diptera amylase, and not targeting amylases from plants or microorganisms present in the substrate. Clear bands were detected in homogenates of feeding substrate from D04 and D07, indicating that BSFL amylase is present in the frass. However, a blurred smear from 65 to 30 kDa with no clear band was obtained with D11 frass homogenate. The second Western blot performed on different homogenates of larvae and substrates from CF gave similar results (Supplementary Figure S5).

Discussion

Growth performance, larval amylase activity and starch digestion

Both in terms of maximal individual fresh weight and FCR, BSFL performance was the best with CF, followed by WB, CGF, WDG and DP. The maximal individual fresh weight of BSFL fed CF (203.7 ± 6.1 mg) was consistent with the literature (Eggink et al., 2023). Chicken feed had a protein+carbohydrate content of 61.6% and a protein:carbohydrate ratio of 1:2, which is close to the nutritional profile recommended for larval growth (40-50% and 1:2-1:4), while WB and CGF deviate from this optimum (Barragan-Fonseca et al., 2021; Cheon et al., 2021). In WDG, limited larval weight gain and high FCR could be caused both by protein excess and low ED of starch (43.1 ± 1.0%). Low larval performance on DP is likely due to its nutritional profile strongly biased towards carbohydrates, but could also result from the presence of alkaloids such as solanine, which are common in potatoes and known to negatively impact the growth of several insect species (Chowanski et al., 2016). Cooking has been identified as a useful pretreatment to improve BSFL and conventional livestock performance on potato-based feed (Ncobela et al., 2017; Moyet et al., 2023).

In CF, CGF and WB, there was no weight gain between D07 and D11, and larval amylase activity peaked on D07 and dropped from D07 to D11. On the other hand, both larval individual weight and amylase activity increased in larvae fed WDG and DP from D04 to D11. We also noticed that about 50%, 35% and 5% of BSFL had reached the dark prepupal stage in CF, WB and CGF respectively (visual rough estimation), while no prepupae were observed in trays with WDG and DP. This suggests that BSFL amylase activity increases with larval growth and drops when larvae stop feeding and prepare for pupation. This pattern has been described in digestive enzymes of insect larvae such as *Ceratitis capitata* (disaccharidase and protease) (Silva et al., 2006), *Heliothis zea* and *Spodoptera exigua* (trypsin) (Broadway and Duffey, 1986), *Spodoptera frugiperda* (amylase, trypsin, lipase) (Lwalaba et al., 2010) or *Rhynchophorus ferrugineus* (protease) (Alarcón et al., 2002) but only hinted in BSFL
so far (amylase, cellulase and protease) (Intayung et al., 2021). This shift of digestive enzymes activity is related to the end of the digestive function of the gut in the preparation of the non-feeding pupal stage, and to the reallocation of resources of the insect from digestion activity to the preparation of metamorphosis (Nestel et al., 2016). Future studies should investigate site-specific variations of amylase activity by separately dissecting larval salivary glands and portions of the gut.

Although larval amylase activity varied with the feeding substrate, no correlation was found between larval amylase activity and the initial substrate nutritional parameters, notably with starch and sugar contents. Based on trials on BSFL fed either a standard diet or a vegetable mix diet with different starch and sugar contents, Bonelli et al., 2020 hypothesized that there could be a positive effect of dietary starch on amylase production (starch induction), or a negative correlation with sugar content (glucose repression). However, this assumption was only based on two feeding substrates and is not supported by the results of the present study (five substrates) nor by the ones of Chen et al., 2023 (four substrates consisting of wheat bran and sawdust added with starch, casein or soybean oil). Yet, a negative correlation between larval amylase activity and reducing sugar content of the larvae was identified. Substrate starch and sugar contents have inconsistent effects on insect amylase activity depending on the species, as reviewed by Bonelli et al., 2020: glucose repression and starch induction have been documented in *Drosophila melanogaster* larvae (Hickey and Benkel, 1982; Inomata et al., 1995), while glucose has a positive effect on amylase activity and starch has none in *Spodoptera frugiperda* larvae (Lwalaba et al., 2010). Whether the observed glucose repression of amylase activity in BSFL results from downregulation of amylase genes expression (Benkel and Hickey, 1986), enzymatic inhibition (Vernon-Carter et al., 2019) or both, remains to be assessed. Interestingly, larval amylase activity was not correlated with initial substrate sugar content, nor with the reducing sugar content of feeding substrate. This inconsistency is likely due to the use of raw agricultural by-products as feeding substrates. Besides starch and sugar contents, these practical diets likely had several parameters potentially affecting amylase activity varying as covariables, such as amino acids profile, presence of amylase inhibitors or inactivation of such inhibitors during the process of each byproduct (Rekha and Padmaja, 2002; Choi et al., 2019), and microbial community (Chen et al., 2023; Meng et al., 2023). To confirm glucose repression in BSFL, further investigation using artificial diets with starch and sugar contents varying independently is required. Characterisation of amylase inhibitors in the substrates would also offer a deeper understanding on the mechanisms of starch digestion (Kaur et al., 2014; Granados-Guzmán et al., 2022).

In fermenting trays, ED of DM and ED of starch varied between substrates but was far from null, which indicates substrate degradation due to microbial activity. Microbial digestion has already been observed in digestibility assays without larvae (Guillaume et al., 2023) and is supported by the fact that we noticed fungi development on the surface of fermenting trays. Significantly high ED of starch both with and without larvae in CGF (100.0 ± 0.0% and 94.0 ± 3.3% respectively) could be due to the relatively low starch content of this substrate that can quickly be degraded, but it also suggests that starch from CGF was highly digestible both for BSFL and microbes. Similarly, WB also had a low starch content and was highly digested by BSFL (100.0 ± 0.0%). However, ED of starch in WB fermenting trays was low (26.8 ± 1.9%), suggesting either that
microbial strains developing on WB were less efficient at digesting starch than those of CGF, or that BSFL amylase is significantly more effective on wheat starch than microbial amylase. These in vivo observations are consistent with the relatively high in vitro amylase activity observed on raw corn and wheat starch (0.27 ± 0.01 U and 0.24 ± 0.01 U respectively). Surprisingly and although it also contained starch from wheat, WDG had a much lower ED of starch than WB, both in feeding and fermenting trays (43.1 ± 1.0% and 6.5 ± 3.2% respectively). Larval growth and ED are two entwined indicators, as bigger larvae ingest more substrate and therefore tend to digest a higher quantity of it, while a more digestible feed is usually associated with improved growth (Waldbauer, 1968; van Loon, 1991; Parra et al., 2012). Consequently, it is also noticeable that, although larvae fed WDG had a similar weight performance than larvae reared on DP, the ED of starch was much higher in DP than in WDG. A potential interpretation could be that, as WDG is a by-product of the wheat distillery process, a significant portion of the original starch had undergone hydrolysis and fermentation to yield ethanol, thereby resulting in the predominance of resistant starch in WDG. Low ED of starch could also be due to the presence of retrograded starch formed during the cooling of cooked wheat grain (Li et al., 2014). High ED of starch observed in feeding trays with DP contradicts the low degradation observed in in vitro assays with raw potato starch. This discrepancy may be attributed to compensatory feeding behaviour: since DP has a relatively low protein content compared to its carbohydrate content, larvae might consume larger quantities of substrate to meet their protein requirements (Le Gall and Behmer, 2014), ultimately resulting in a higher amount of starch digested. Substrates ranked differently in terms of maximal larval amylase activity and ED of starch, highlighting that amylase activity in BSFL was not a good proxy of starch digestibility.

In vitro raw starch digestion assays indicated that BSFL amylase was 13 times more efficient on starch from rice and 9 times more on starch from corn or wheat, than on potato starch. Existing literature supports the greater in vitro and in vivo digestibility of cereal-derived starch compared with potato starch (Dreher et al., 1984). The reported difference in starch granules diameter highlighted that granule size is a major driver of degradability as, for the same starch quantity, smaller granules offer a higher surface area for interaction with amylases than bigger granules (Svihus et al., 2005; Jagadeesan et al., 2020). Reported starch granules sizes were consistent with the literature (Pérez and Bertoft, 2010; Jagadeesan et al., 2020; Junejo et al., 2022). We observed that granules had a consistent size between raw starches and feeding substrates of the same plant origin, except for DP starch granules that were bigger (up to 150 µm) than in raw potato starch (30-50 µm), presumably because of potato strain differences (de Pater et al., 2006). Amylose content was not measured in our study and varies within a plant species (Junejo et al., 2022) but could also explain degradability differences of raw starches, as high amylose content has been associated with impaired starch digestion (Jagadeesan et al., 2020). Finally, ultrastructural aspects of starch granules (e.g. crystalline or amorphous structure, degree of polymerisation, number of branched chains, chains length) can also affect starch resistance regarding hydrolysis by amylases (Pérez and Bertoft, 2010; Junejo et al., 2022). In feeding substrates, physical protection of starch granules by cell walls and starch retrogradation after cooking and cooling can also lead to resistant starch (Junejo et al., 2022). These observations highlight the importance of considering
digestibility and not only crude starch content when formulating an optimised diet for
BSFL.

Larval amylase optimal pH and temperature

The optimal pH range of 5.0-6.5 was consistent with the observations of Bonelli et al., 2019 and aligns with the pH of 6.0 observed in the anterior midgut lumen of BSFL, the gut region where amylases are secreted by columnar cells. However, we observed a higher amylase activity in acidic conditions (pH 4.0-5.5) and a lower activity in alkaline conditions (pH 7.0-8.5) than in this previous study. This is likely due to differences in sample preparation, as we worked with purified amylase from whole larvae homogenate and not midgut juice, or to different black soldier fly strains and rearing conditions. Surprisingly, Intayung et al., 2021 reported that amylase activity of whole ground BSFL was slightly decreasing for pH varying between 3.0 and 9.0, instead of displaying the usual bell-shape trend, which was not commented on by the authors. Amylase of BSFL has a lower optimal pH than in most other Diptera species (pH 7.0-8.5) (Da Lage, 2018), except for the generalist saprophagous larvae of *Musca domestica* (pH 6.5) (Terra and Ferreira, 1994). This could be an adaptation of *H. illucens* and *M. domestica* larvae to efficiently digest carbohydrates from acidic fermenting substrates.

We observed that purified BSFL amylase operates on a large range of temperatures. The optimum of 70°C that we identified was higher than the one reported by Intayung et al., 2021 on unpurified larvae homogenate (55-60°C depending on larval age). This is likely related with different sample preparation, as it has been reported that the curve of amylase activity with temperature of whole adult *D. melanogaster* homogenate was flat between 30 and 45°C (Shibata and Yamazaki, 1994), while for purified amylase, the curve was sharper and peaked at 52°C (Cipolla et al., 2012). The 70°C optimal temperature of BSFL amylase was higher than the optimum of amylase from other Diptera such as *M. domestica* and *Sarcophaga ruficornis* (50°C) (Sinha, 1975) or *D. melanogaster* (52°C) (Cipolla et al., 2012) and from other insects (35-60°C) (Kaur et al., 2014). While 70°C is above estimations of the maximal lethal temperature for BSFL (35-44°C) (Chia et al., 2018; Wang et al., 2023), biologically realistic conditions around 35°C still allow to reach 40% of maximal amylase activity. Higher temperatures up to 40-45°C can be reached and could foster starch digestion through social feeding when larvae aggregate and generate heat due to their movement (Parra Paz et al., 2015; Li et al., 2023). Comparatively, the optimal temperature of BSFL proteases has been measured around 45°C (Kim et al., 2011; Bonelli et al., 2019).

Nine copies of genes coding for alpha-amylase-like-proteins have been identified in the *H. illucens* genome (NCBI accession number GCF_905115235.1): consequently, we likely observed the overall activity of a mix of different amylase isozymes, which could differ in their optimal conditions, but also in their substrate specificity or regions of expression (Kim et al., 2011). Precise characterisation of each amylase isozyme by genetic engineering could provide insight into their functions and possible complementarity.
## Substrate amylase activity, Western blot and external digestion

Maximal amylase activity was identified in initial substrates, before inoculation with starter larvae. This was likely due to amylases from the plant materials of each substrate, as grains and tubers contain alpha- and beta-amylases (Sundarram and Murthy, 2014; Guzmán-Ortiz et al., 2018). Differences between treatments (i.e. substrate from fermenting or feeding tray, and sampling date) were only detected in CF, CGF and WB. For each of these three substrates, amylase activity without larvae varied with time following a different pattern, but activity on D11 was always lower than on D00. It is to be noticed that we visually observed significant fungi development on the surface of each fermenting substrate and, although fungi and bacteria can produce amylase to digest starch (Farooq et al., 2021), no increase of amylase activity in fermenting substrates was detected. We assume that vegetal amylases were degraded during the 11 days of fermentation, possibly used as a protein source by the microorganisms developing in the substrates. In CF, CGF and WB, the amylase activity of substrate in feeding trays decreased with time and the lowest activity of all conditions was reached on D11 in feeding trays. The decrease of amylase activity over time in both fermenting and feeding trays contrasts with the increase in catalase, urease, sucrase, protease, cellulase and lipase activity described by Tao et al., 2023 in a fermented food waste substrate both with and without BSFL inoculation. The authors also reported that all enzymatic activities were higher in substrates with BSFL compared to fermenting substrates, suggesting that BSFL release digestive enzymes in the substrate or have a positive effect on microbes releasing enzymes, which is inconsistent with our results. This discrepancy could be explained by a higher microbial charge in food waste, resulting both from the nature of the substrate and from the application of a preliminary fermenting step (Tao et al., 2023). Our data also contrast with the increase of substrate amylase activity over time observed with *A. ludens* larvae, while no variation occurred without larvae (Aceituno-Medina et al., 2023). Our results suggest that either BSFL and their microbiota degrade initial vegetal amylases and that larvae do not perform external digestion, or that BSFL do release amylase in their substrate, but to a much lesser extent than the degradation rate of plant amylase, resulting in an overall decrease of measured amylase activity. No significant difference in substrate amylase activity was detected between treatments in WDG and DP, because both soluble protein content and maltose release in the presence of the starch solution were low and close to the detection limit in these homogenates. As a result, computed amylase activity values were floating with no significant differences between treatments. Assays on substrate homogenates also revealed that amylase activity was much lower in WDG and CGF than in DP, CF and WB. This could be explained by the different production processes of each substrate. Indeed, while WDG results from wheat grain that has been cooked and fermented (Heuzé et al., 2017) and CGF went through a high-temperature sulfur-dioxide steep (Heuzé et al., 2015), DP, CF and WB have not gone through any strong processing step that could potentially deteriorate or inactivate vegetal amylases.

No correlation was found between initial substrate nutritional parameters and both feeding and fermenting substrate amylase activity, suggesting that amylase activity in substrates is a phenomenon intricately related to substrate characteristics. Microbial
communities of the substrates were not investigated but were likely involved in the different patterns of substrate amylase activity observed (Chen et al., 2023). Amylase activity in feeding substrates was neither correlated with amylase activity in larvae, nor with that in fermenting substrates. We concluded that the amylase activity of the three types of samples have different evolution patterns.

Western blot results indicate that BSFL amylases of the same molecular weight were detected in larvae and frass homogenates, but not in native or fermented substrates. These observations support the hypothesis that amylases from BSFL are present in consumed substrates, but are associated with a marginal additional amylase activity compared to the reduction of native vegetal amylase due to larval and microbial digestion. Both amylases from BSFL and *D. melanogaster* were detected at the same weight of 44 kDa, which is lower than the value of 52 kDa reported in the literature based on the amino acid sequence of *D. melanogaster* amylase (Da Lage, 2018). As a molecular weight observed on a Western blot is not as precise as a calculated value (Kroon et al., 2022), BSFL amylase should also be considered to have a weight around 52 kDa. Secondary bands that were detected with larvae homogenates suggest that at least three isozymes could be expressed successively during BSFL development, with a molecular weight of 41 kDa (from D04 to D07), 52 kDa (during overall larval development) and 59 kDa (from D07 to D11) respectively. These faint bands could also result from post-traductional modifications or partial breakdown of native amylase by protease. Nevertheless, only 52 kDa amylase was reported in consumed substrate homogenates, possibly indicating that only one of the isozymes was released in the substrate by BSFL. The smear that we noticed with D11 consumed substrate homogenate did not allow for proper amylase identification. As it was also observed with a duplicate sample in the second Western blot, it is likely not due to a mistake from the operator, but probably results from improper protein migration due to alkaline conditions in late frass homogenates (Meneguz et al., 2018). The imager was still able to detect a signal, as the Western blot result from D11 frass homogenate is unambiguously different from what was observed with native or fermented substrates. Therefore, although we cannot clearly identify the molecular weight of the amylase in this sample, we are confident that BSFL amylase was present in the feeding substrate on D11. To further confirm that BSFL release amylase in their feeding substrate, several strategies could be proposed, such as observing the color of agar cubes containing starch and iodine in the presence of BSFL (Sakaguchi and Suzuki, 2013), or measuring amylase activity in an artificial sterile amylase-free substrate after consumption by BSFL. Amylases in frass samples can either come from (a) active release from the salivary glands of BSFL, (b) passive release through BSFL faeces or (c) passive release from potential decaying dead BSFL in the substrate, although none were observed during the sampling. More advanced feed trial designs should be developed to identify the mechanisms behind the liberation of these enzymes, such as using a mean of individual BSFL immobilisation to separately collect substrate in contact with mouthpieces and faeces, and assess amylase activity in both fractions. Furthermore, as BSFL increase the pH of their substrate up to 9.0 during bioconversion (Ma et al., 2018; Meneguz et al., 2018), additional investigations are needed to determine if larval amylases are active in the substrate and to explore their role in external digestion. Performing Western blots with antibodies targeting plant and microbial amylases would also allow to confirm that the amylase activity measured in substrates without larvae was due to native amylase and microbes.
**Conclusion**

This study is the first one to report that BSFL amylase activity increases during larval growth and drops when approaching pupation. It is also the first to provide optimal pH (5.0-6.5) and temperature (70°C) for purified amylase of BSFL. Variations of amylase activity in larvae were observed between different feeding substrates but could not be correlated with their nutritional parameters. However, we identified that larval reducing sugar content and amylase activity were negatively correlated, suggesting glucose repression of amylase production in BSFL. Our results indicated that BSFL amylase is more efficient on rice starch than on corn and wheat starch, and can hardly digest potato starch. Enzymatic efficiency differences partly explained variations in FCR and ED between substrates, hinting at more complex feeding mechanisms such as compensatory feeding and insect-microbes interaction. Amylases from BSFL were found in their substrate after feeding, a first argument in favour of external digestion in BSFL, but there was no associated increased amylase activity in the substrate. Further work is required on the regulation of amylase production both by larvae and their microbiota, and to explore the mechanisms of digestive enzymes released in the feeding substrate by BSFL as well as their role in external digestion.

**Ethics approval**

Not applicable.

**Data and model availability statement**

None of the data were deposited in an official repository, but they are available from the corresponding authors upon request. The R code used to analyse the data is available in the Supplementary Material S1.

**Declaration of Generative AI and AI-assisted technologies in the writing process**

The authors did not use any artificial intelligence assisted technologies in the writing process.
**Author ORCIDS**

J.B. Guillaume: https://orcid.org/0000-0003-3678-3855

J.-L. Da Lage: https://orcid.org/0000-0002-1372-6762

S. Mezdour: https://orcid.org/0000-0002-5191-0421

F. Marion-Poll: https://orcid.org/0000-0001-6824-0180

C.M.C. Brouzes: https://orcid.org/0000-0001-6433-901X

P. Schmidely: https://orcid.org/0000-0002-7566-7868

**Declaration of interest**

This work was funded by Agronutris, a company producing insect-based ingredients for animal feed. The company did not interfere with the conclusions of the study.

**Acknowledgements**

Authors would like to thank Daniele Bruno (University of Insubria), Marie Merle (Sorbonne Université), Gaëlle Claisse and Samuel Gornard (IDEEV) for their technical support and advice on molecular biology assays, along with the R&D Team of Agronutris for their inputs on the experimental approach and for providing the starter larvae.

**Financial support statement**

This work was funded by Agronutris, with support from the French Ministry of Higher Education, Research and Innovation (CIFRE grant n°2019-1881 attributed to J.B.G.).
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# Tables

<caption>**Table 1.**</caption>

Nutritional parameters of the feeding substrates for black soldier fly larvae.

<table><thead><tr><th>Item</th><th>CF</th><th>CGF</th><th>WB</th><th>WDG</th><th>DP</th></tr></thead><tbody><tr><td>Starch (% DM)</td><td>34.6</td><td>11.1</td><td>12.5</td><td>26.1</td><td>47.5</td></tr><tr><td>Sugars (% DM)</td><td>4.8</td><td>0.8</td><td>5.3</td><td>4.1</td><td>8.4</td></tr><tr><td>Carbohydrate (% DM)</td><td>39.4</td><td>11.9</td><td>17.8</td><td>30.2</td><td>55.9</td></tr><tr><td>Protein (% DM)</td><td>22.2</td><td>22.3</td><td>17.6</td><td>24.2</td><td>9.6</td></tr><tr><td>Protein+carbohydrate (% DM)</td><td>61.6</td><td>34.3</td><td>35.5</td><td>54.4</td><td>65.5</td></tr><tr><td>Protein:carbohydrate</td><td>1:2</td><td>2:1</td><td>1:1</td><td>1:1.2</td><td>1:6</td></tr></tbody></table>

Abbreviations: CF = chicken feed; CGF = corn gluten feed; WB = wheat bran; WDG = wheat distillers grain; DP = discarded potatoes.
**Table 2.**

Description of the microscopic aspect of starch granules from substrates used as feed for black soldier fly larvae and raw starches.

<table><thead><tr><th rowspan="2">Item</th><th colspan="5">Substrates</th><th colspan="4">Raw starches</th></tr><tr><th>CF</th><th>CGF</th><th>WB</th><th>WDG</th><th>DP</th><th>Corn</th><th>Wheat</th><th>Potato</th><th>Rice</th></tr></thead><tbody><tr><td>Diameter (µm)</td><td>5-30</td><td>10-20</td><td>10-30</td><td>5-30</td><td>50-150</td><td>10-20</td><td>10-20</td><td>30-50</td><td>&lt; 5</td></tr><tr><td>Shape</td><td>Round</td><td>Angular</td><td>Round</td><td>Round</td><td>Oval</td><td>Angular</td><td>Round</td><td>Oval</td><td>Angular, tends to agglomerate</td></tr></tbody></table>

Abbreviations: CF = chicken feed; CGF = corn gluten feed; WB = wheat bran; WDG = wheat distillers grain; DP = discarded potatoes.
## Figure captions

**Fig. 1.**

Experimental design of the study. Number of replicates (n) are for each substrate. CF: chicken feed; CGF: corn gluten feed; WB: wheat bran; WDG: wheat distillers grain; DP: discarded potatoes; D00, D04, D07 and D11: start of the experiment, fourth, seventh and eleventh days of the experiment, respectively; '+' and '-' indicate presence or absence of black soldier fly larvae in the tray (feeding or fermentation tray) respectively. ED of DM: estimated DM digestibility; FCR: feed conversion ratio.

**Fig. 2.**

Effect of substrate (S) and time (T) on black soldier fly larvae individual fresh weight. CF: chicken feed; CGF: corn gluten feed; WB: wheat bran; WDG: wheat distillers grain; DP: discarded potatoes; D04, D07 and D11: four, seven and eleven days of feeding, respectively. Values are reported as mean ± SEM of 30 individuals. Different letters indicate significantly different values (generalised linear model with gamma distribution, post-hoc pairwise comparison with Bonferroni adjustment, threshold: p-value<0.05).

**Fig. 3.**

Effect of substrate on feed conversion ratio (on a dry basis) of black soldier fly larvae after 11 days of feeding. CF: chicken feed; CGF: corn gluten feed; WB: wheat bran; WDG: wheat distillers grain; DP: discarded potatoes. Values are reported as mean ± SEM of three feeding trays. Different letters indicate significantly different values (generalised linear model with gamma distribution, post-hoc pairwise comparison with Bonferroni adjustment, threshold: p-value<0.05).

**Fig. 4.**

Effect of substrate (S) and presence of black soldier fly larvae (L) on estimated digestibility of (A) DM and (B) starch after 11 days. CF: chicken feed; CGF: corn gluten feed; WB: wheat bran; WDG: wheat distillers grain; DP: discarded potatoes; '+' and '-' indicate the presence or absence of larvae in the tray (feeding or fermenting tray) respectively. Values are reported as mean ± SEM of three trays. Different letters indicate significantly different values (generalised linear model with Gaussian distribution, post-hoc pairwise comparison with Bonferroni adjustment, threshold: p-value<0.05).
Fig. 5.

Effect of (A) pH and (B) temperature on amylase activity of purified amylase from black soldier fly larvae. Values are reported as mean ± SEM of three to six technical replicates. Values significantly different from the maximal amylase activity (A: activity at pH 6.5; B: activity at 70°C) are indicated by asterisks (generalised linear model with gamma and Gaussian distribution respectively, post-hoc Dunnett comparison, threshold: p-value<0.05).

Fig. 6.

Effect of substrate (S) and time (T) on black soldier fly larvae amylase activity. CF: chicken feed; CGF: corn gluten feed; WB: wheat bran; WDG: wheat distillers grain; DP: discarded potatoes; D04, D07 and D11: four, seven and eleven days of feeding, respectively. Values are reported as mean ± SEM of three homogenates prepared with at least 10 larvae. Different letters indicate significantly different values (generalised linear model with gamma distribution, post-hoc pairwise comparison with Bonferroni adjustment, threshold: p-value<0.05).

Fig. 7.

Relation between sample reducing sugar content and amylase activity in black soldier fly larvae homogenates. N=48 (5 substrates × 3 sampling dates × 3 homogenates + 3 starter larvae samples).

Fig. 8.

Effect of presence of black soldier fly larvae (L) and time (T) on amylase activity of each substrate: (A) chicken feed (CF), (B) corn gluten feed (CGF), (C) wheat bran (WB), (D) wheat distillers grain (WDG) and (E) discarded potatoes (DP). '+' and '-' indicate the presence or absence of larvae in the tray (feeding or fermenting tray) respectively. D00, D04, D07 and D11: initial substrate, substrate after four, seven and eleven days of the experiment, respectively. Values are reported as mean ± SEM of three homogenates prepared with 1.0 g of substrate. Different letters indicate significantly different values (generalised linear model with Gaussian distribution, post-hoc pairwise comparison with Bonferroni adjustment, threshold: p-value<0.05).

Fig. 9.

Hydrolysis of different types of raw starch by purified black soldier fly larvae amylase. Values are reported as mean ± SEM of four technical replicates performed with the same larvae homogenate. Different letters indicate significantly different values
(generalised linear model with Gaussian distribution, post-hoc pairwise comparison with Bonferroni adjustment, threshold: p-value<0.05).

Fig. 10.

Detection of amylase in black soldier fly larvae and substrate homogenates by Western blot with a primary antibody targeted against *Drosophila melanogaster* amylase. MW: molecular weight; L: homogenate of larvae fed chicken feed; CF- and CF+: chicken feed substrate from fermenting tray and feeding tray, respectively; D00, D04, D07 and D11: larvae or substrates were sampled at the start or after four, seven and eleven days of experiment respectively. Control: purified *D. melanogaster* amylase. Blue lines and pink horizontal lines indicate the outline of each lane and the identified bands, respectively, which are automatically detected by the Image Lab software.
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A

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B

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<figure><img src="image_13.png" /><figcaption>Amylase activity in substrates (µmol maltose/s/g protein)</figcaption></figure>
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