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Article

Investigation of the Phageome and Prophages in French Cider, a Fermented Beverage

by
Pierre Ledormand
1,
Nathalie Desmasures
1,
Cédric Midoux
2,3,4,
Olivier Rué
2,3 and
Marion Dalmasso
1,*
1
Normandie Univ., UNICAEN, UNIROUEN, ABTE, 14000 Caen, France
2
INRAE, MaIAGE, Université Paris-Saclay, 78350 Jouy-en-Josas, France
3
INRAE, BioinfOmics, MIGALE Bioinformatics Facility, Université Paris-Saclay, 78350 Jouy-en-Josas, France
4
INRAE, PROSE, Université Paris-Saclay, 92761 Antony, France
*
Author to whom correspondence should be addressed.
Microorganisms 2022, 10(6), 1203; https://doi.org/10.3390/microorganisms10061203
Submission received: 24 May 2022 / Revised: 1 June 2022 / Accepted: 8 June 2022 / Published: 12 June 2022
(This article belongs to the Special Issue Phages from the Fermented Products)
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Abstract

:
Phageomes are known to play a key role in the functioning of their associated microbial communities. The phageomes of fermented foods have not been studied thoroughly in fermented foods yet, and even less in fermented beverages. Two approaches were employed to investigate the presence of phages in cider, a fermented beverage made from apple, during a fermentation process of two cider tanks, one from an industrial producer and one from a hand-crafted producer. The phageome (free lytic phages) was explored in cider samples with several methodological developments for total phage DNA extraction, along with single phage isolation. Concentration methods, such as tangential flow filtration, flocculation and classical phage concentration methods, were employed and tested to extract free phage particles from cider. This part of the work revealed a very low occurrence of free lytic phage particles in cider. In parallel, a prophage investigation during the fermentation process was also performed using a metagenomic approach on the total bacterial genomic DNA. Prophages in bacterial metagenomes in the two cider tanks seemed also to occur in low abundance, as a total of 1174 putative prophages were identified in the two tanks overtime, and only two complete prophages were revealed. Prophage occurrence was greater at the industrial producer than at the hand-crafted producer, and different dynamics of prophage trends were also observed during fermentation. This is the first report dealing with the investigation of the phageome and of prophages throughout a fermentation process of a fermented beverage.

1. Introduction

Bacteriophages (phages) are viruses infecting bacteria, and that can establish lytic, chronic or temperate (lysogenic) infections. To designate phage communities, the term “phageome” is frequently used in the scientific literature. However, it seems that no consensual definition of this term has been retained to date. In some cases, the term “phageome” seems to encompass communities of free lytic phages and of prophages (temperate phages included in bacterial genomes) [1]. Nevertheless, the majority of the studies dealing with phageomes often consider free lytic phages, without including prophages [2,3]. Consequently, the term “phageome” will be used in the current study for designating free lytic phage communities only. Prophages will be considered separately. Phageomes are studied in multiple environments such as the human gut [1,2,3], the human oral cavity [4], plants [5], oceans [6], and more recently, glaciers [7]. Studies of the phageomes in fermented foods and beverages are scarce [8]. To date, only three studies strictly describing the phageomes of Asian fermented food products such as kimchi, sauerkrauts and fermented shrimps [9,10], and of a French cheese [11] are available, but none about fermented beverages. Environmental sources of phages in cheese have also been studied by investigating the airborne viromes (whole virus communities) in two cheese production plants [12]. Prophages (temperate phages) are also well known for their impact on the functioning of the bacterial cells that carry them, either through beneficial or non-beneficial interactions [13]. Their impact at the scale of a microbial community remains underexplored, and still needs to be taken into account [14]. In this way, exploring the diversity of prophages also seems essential to assess their role in microbial fluxes within ecosystems such as fermented foods and beverages, as phages are recognised to be the drivers of microbial diversity [15].
Although the phageomes of fermented foods are not well described yet, their presence in such matrices has been known for decades. Lytic phages are extensively studied for their antibacterial properties against pathogenic bacteria encountered in the food industry such as, for example, Salmonella sp. [16] and Listeria monocytogenes [17], and against spoilage bacteria such as Acetobacter in wine [18] and Levilactobacillus brevis in beer [19]. Lytic phages are also known as killers of such industrial starters as in dairy products, where phages targeting Lactococcus lactis and Streptococcus thermophilus [20] can cause fermentation defects. No study is available about the phageomes in fermented beverages, especially in cider. Cider is an alcoholic fermented apple beverage, consumed all around the world, presenting harsh physico-chemical conditions with low pH values (3.0–4.0) and the presence of ethanol [21], polyphenols and volatiles [22]. Two types of fermentation occur during cider elaboration. First, the alcoholic fermentation is performed by yeasts, including the Saccharomyces species (S. cerevisiae and S. uvarum) and non-Saccharomyces species (Hanseniaspora, Candida) [21]. Then, the malolactic fermentation (MLF) is performed by Lactic Acid Bacteria (LAB) belonging to Oenococcus sp., Leuconostoc sp., Pediococcus sp. and Lactobacillus sensu lato [22]. Describing phageome diversity in cider is thus interesting to better understand the microbial dynamics during the fermentation processes. The aim of this work was to study the diversity of the phageome, and also the prophage distribution in cider, throughout the fermentation process in two cider tanks from two producers. Several methods were thus tested to extract free phage particles from cider, in order to explore the phageome, and prophages were searched for inside bacterial metagenomes throughout the cider fermentation.

2. Materials and Methods

2.1. Cider Sample Collection for Phageome and Prophage Investigation

A tank from an industrial producer (B) and a tank from a hand-crafted producer (P), located in the Normandy region (France), were sampled during the first month of the cider fermentation process, at four time points, to explore the presence of free phage particles and of prophages. Details about the collected samples are given in Table 1.

2.2. Methodological Developments for Collecting Phage Particles from Cider and for Phageome DNA Extraction

To collect and extract total phage DNA from cider, different approaches were tested and are summarised in Table 2.

2.2.1. Validation of Methods on Cider Samples Spiked with Phages

First, phageome DNA extraction was tested on cider samples spiked with phage UCMA 21115, previously isolated from cider [23] and originating from the UCMA collection (Université de Caen Microbiologie Alimentaire, Caen, France), in order to validate the method used. Concentrations of phage UCMA 21115 ranging from 104 to 109 PFU/mL were used (Table 2—I, II and III). Spiked cider samples (50 mL) were centrifuged at 4700× g for 20 min at 4 °C, and the pellet was suspended into 500 µL SM buffer (200 mM NaCl, 10 mM MgSO4, 50 mM Tris pH 7.5) (Table 2—I). Phage precipitation with PEG-8000 (w/v) and 0.5 M NaCl was also tested by adding them directly to spiked cider, adjusted (Table 2—III) or not (Table 2—II) to pH 5.5. PEG-8000 addition was followed by an incubation step at 4 °C overnight to maximise the precipitation of phage particles. Then, samples were centrifuged, as described above, and the pellets were suspended into 500 µL SM buffer.
In all cases, one volume of chloroform was added to the pellets or to the precipitates suspended into 500 µL SM buffer beforehand, in order to eliminate membrane vesicles [11]. Samples were then centrifuged at 8000× g for 5 min. The aqueous phase was treated with DNAse (Fisher Bioreagents, Vilnius, Lithuania) and RNAse (Thermofisher, Vilnius, Lithuania) for 1 h at 37 °C to eliminate non-viral DNA. Then, 10% SDS and 20 mg/mL proteinase K (Thermofisher, Vilnius, Lithuania) were added and an incubation step at 56 °C for 20 min was performed [24]. Finally, phage DNA were extracted with three different methods. The first one (named buffer 1), based on a method described for faecal samples, consisted in the addition of 100 µL Phage Lysis Buffer (4.5 M guanidinium isothiocyanate, 44 mM sodium citrate pH 7.0, 0.88% sarkosyl, 0.72% 2-mercaptoethanol) [2]. In the two other methods, Phage Lysis Buffer was supplemented with 1% (buffer 2) or 2% (buffer 3) of polyvinylpolypyrrolidone (PVPP), in order to counteract the effects of cider polyphenols. These approaches were based on previous protocols designed for plant DNA extraction [25]. Samples were then incubated at 65 °C for 10 min.
Phage DNA purification was performed by two successive steps of phenol:chloroform:isoamyl 25:24:1 (Sigma Aldrich, St. Louis, MO, USA) (v/v) treatment and centrifugation at 8000× g for 5 min. DNA precipitation was performed with 2.5 volumes of absolute ice-cold ethanol before a centrifugation step at 13,000× g for 15 min at 4 °C. DNA was eluted in 50 µL of water, and quantified using a Nano TECAN spectrophotometer (Life Science, Zürich, Switzerland). DNA integrity was checked on a 1% agarose gel.

2.2.2. Application to Non-Spiked Cider Samples

Phageome DNA extraction protocols were then applied to cider samples (not spiked with exogenous phage UCMA 21115), using different volumes (20, 50 and 1000 mL) (Table 2—IV, V, VI, VII). Cider samples were centrifuged at 4700× g for 20 min at 4 °C, and DNA extractions were performed on the pellets suspended into 500 µL SM buffer beforehand (Table 2—IV) and on the supernatants (Table 2—V) to maximise the chances of obtaining phage DNA. Supernatants were first filtered through 0.45 µm filters and precipitated overnight with PEG-8000 at 4 °C, before being centrifuged at 4700× g for 20 min at 4 °C. The resulting pellets were suspended into 500 µL SM buffer prior to DNA extraction, as described above.
DNA extractions were also directly performed after the addition of PEG-8000 and NaCl, prior to the first centrifugation step, on cider samples adjusted to pH 5.5 (Table 2—VII) or not (Table 2—VI). The precipitates were centrifuged as described above, and the pellets were suspended into 500 µL SM buffer. DNA was extracted following the same methods as described above.

2.2.3. Optimisation of the Method to Collect Phage Particles from Cider

To concentrate phage particles from cider, 400 mL of cider was filtered with a tangential flow filtration (TFF) method (Table 2—VIII). The TFF system used was the Vivaflow® 50 (Sartorius, Göttingen, Germany), a PES membrane cassette with a total surface area of 50 cm2 and a molecular weight cut-off of 100 kDa. In order to validate the functioning of the TFF system, cider was spiked with exogenous phage UCMA 21115, at a concentration of 105 PFU/mL. Retentate was filtered through 0.45 µm filters before DNA extraction, as described above.
The second method used to concentrate phage particles was the chemical Fe-based flocculation, as described for the concentration of ocean viruses from large volume samples (Table 2—IX) [26]. As for TFF, 400 mL of cider spiked with exogenous phage UCMA 21115, at concentrations ranging from 104 to 108 PFU/mL, was tested. Briefly, FeCl3 was added to cider samples to obtain a final concentration of 1 mg/L before incubation at room temperature for 1 h to allow Fe-phage complexes to flocculate. Then, the solutions were filtered on Whatman polycarbonate membrane filters with 0.8 µm-pores. Finally, filter retentates were suspended into 10 mL suspension buffer (0.25 M ascorbic acid, 0.2 M Mg2EDTA, pH 6.0) before DNA extraction, as described above.
In parallel, phage isolation was also performed on these retentates from TFF and flocculation, potentially enriched in phages. Isolation and propagation were carried out on 160 strains from genera found in cider (Table S1), using the classical double layer plate technique with MRS 0.5% (w/v) supplemented with 10 mM CaCl2 and 10 mM MgSO4, and incubated at 30 °C for 24 h to allow for the formation of plaques.

2.3. Microbial DNA Extraction for Prophage Investigation

At each sampling time (Table 1), 50 mL of cider samples were centrifuged at 4700× g for 20 min at 4 °C. Total DNA from pellets was extracted and purified using the NucleoSpin Soil kit (Macherey Nagel, France) according to the manufacturer’s instructions. DNA quality and quantity were checked as described above.

2.4. DNA Sequencing and Bioinformatics Analysis

DNA library preparation and sequencing were carried out at Genewiz facilities (Leipzig, Germany), on an Illumina HiSeq sequencer producing paired-end reads of 2 × 150 bp in length. To access the quality of raw reads FastQC v.0.11.9 [27] was used. FastP v 0.20.0 [28] was used to eliminate reads with a length inferior to 50 nucleotides and bases with a bad quality (lower than 15). Sequence assembly was performed with SPADES v 3.14.0 with option—meta [29]—and only contigs longer than 1 kb were kept. Moreover, taxonomic assignations to access the composition of trimmed reads was done using kaiju v 1.7.3 [30] against database nr_euk (103 M protein sequences from nr: Bacteria, Archaea, Viruses, Fungi and microbial eukaryotes). The sequencing data from this study are available in NCBI SRA repository, reference number PRJNA803977.

2.5. Prophage Prediction

VIBRANT was used for prophage prediction [31]. VIBRANT is a hybrid machine learning used to maximise identification of lytic viral genomes and integrated proviruses, including highly diverse viruses. VIBRANT was used separately for each sample and contigs identified with VIBRANT were clustered between all samples using cd-hit-est [32] with default parameters (global sequence identity 90%). Reads were finally mapped on this set of contigs identified as potential phages. To generate abundance matrices, mapping results were converted into reads per length of the contig (in kb) per million (RPKM) and RPKM values of contigs above 5.0 were kept for analyses. Contigs identified by VIBRANT were aligned with BLAST against the NCBI non-redundant nucleotide database (E-value < 10−16 and percentage of identity >90%) to generate information about the host.

3. Results

3.1. Development of Methods for Phageome DNA Extraction from Cider

In order to explore the diversity of the phageome of cider, the total phage DNA was intended to be extracted from cider samples. Consequently, several methods for the collection of phages were tested (Table 2). For each type of sample preparation, three DNA extraction methods were performed (buffers 1, 2 and 3).

3.1.1. Validation of the Extraction Method after Collection of Phage Particles from Spiked Cider Samples

Phageome DNA extraction methods adapted from other matrices [2,11] were first tested on cider samples spiked with phage UCMA 21115 [23], at concentrations ranging from 104 to 109 PFU/mL. In some cases, the pH of the samples was adjusted to 5.5 (Table 2—III–VII), as it corresponded to a pH value suitable for phage DNA extraction from cider in preliminary experiments (data not shown).
This step was also intended to determine the threshold of phages necessary to collect enough DNA in this matrix. In all the tested conditions, phage DNA was only extracted in cider samples adjusted to pH 5.5 after PEG precipitation (Table 2—III). At phage concentrations inferior to 107 PFU/mL, no phage DNA was detected on agarose gel (Figure S1) whatever the DNA extraction buffer used, and very low quantities of DNA were obtained (less than 5–10 ng/µL). Phage DNA was detected at phage concentrations starting from 107 PFU/mL whatever the DNA extraction buffer used (Figure S1). No phage DNA was extracted (data not shown) from spiked cider samples when the pH was not adjusted to 5.5 (Table 2—I,II). This result showed that phage DNA extraction methods were only efficient at a phage concentration starting from 107 PFU/mL and at pH 5.5.

3.1.2. Extraction of Phage Particles from Non-Spiked Cider Samples

The methods to collect phages and to extract the phageome DNA were then tested in non-spiked cider samples (Table 2—IV–VII). No phage DNA was extracted regardless of the tested buffers, the sample volume or the sample preparation (pellets, supernatants, cider concentrated with PEG and cider adjusted to pH 5.5 before PEG precipitation) (data not shown). As no phage DNA could be recovered in the non-spiked cider samples, whatever the tested conditions, it suggested a low occurrence of lytic phages in cider (probably less than 107 PFU/mL), resulting in the difficulty to extract phageome DNA in such a fermented beverage.

3.1.3. Viral Concentration Methods Were Tested to Improve the Phageome DNA Extraction

The previous results revealed a presumably low quantity of lytic phages in cider shown by the difficulty to extract phageome DNA from non-spiked cider samples. Thus, concentration methods were used in order to improve the phageome DNA extraction, starting from larger volumes of cider than the volumes tested above.
TFF was intended to be used to concentrate phages from up to 5 L of cider. A primary attempt was performed with a reduced batch of 400 mL of cider spiked with phage UCMA 21115 at a concentration of about 1.9 × 105 PFU/mL (i.e., resulting in a total quantity of 7.6 × 107 phages in 400 mL) as a positive control and to set the filtration parameters. At first, TFF appeared to be a suitable method to concentrate phages in cider samples because the total quantity of spiked phages was recovered in the retentate, with 8 × 107 collected phages compared to the calculated 7.6 × 107 expected phages. Concerning the filtration parameters, the flow rate was of 120 mL per hour for this cider sample, whereas it is rather around 750 mL per hour for sea water samples (supplier data). The attempts to filter (0.45 µm) the retentate (15 mL) to remove yeasts, bacteria and other cider components failed, as the cider retentate was too thick. As a result, phages were lost at this step with only 3.4 × 104 phages that were recovered, which represents a loss of 3 log10. In view of these results, the option to use TFF in larger volumes of non-spiked cider samples was abandoned.
Another method to concentrate phages was tested, the iron (Fe)-based flocculation, which has the advantage of using large-pore-size filters (0.8 µm) compared to TFF. Concerning cider samples spiked with phage UCMA 21115, phage recovery was quite good at high phage concentrations. It was of 8.5 × 108 for cider samples, for an initial inoculation of 4.0 × 1010, representing a phage loss of about ≈1 log10 (Table 3). At low phage concentrations, the phage recovery with Fe-based flocculation was less efficient. For example, with an initial inoculation of 4.0 × 106, 5.0 × 103 phage particles were retrieved in cider samples (Table 3). In addition, filtration of the cider sample was also quite complicated due to clogging. In view of these results, the option to use (Fe)-based flocculation on larger volumes of non-spiked cider samples was also abandoned.
In addition, the isolation of phages was attempted by using 160 potential host strains from genera found in cider, from the TFF retentate and after cider flocculation, to check the presence of potential lytic phages on these bacteria. No phage, except phage UCMA 21115 used as positive control, was retrieved from these samples, confirming that the phageome of cider was not abundant, and thus not easily accessible.

3.2. Prophage Investigation in Two Cider Tanks during Cider Fermentation

To complete the data concerning the lytic phageome, prophages present inside the metagenomes of the bacterial community involved in the cider fermentation process were investigated. As described in Table 1, cider sampling was carried out during the first month of cider fermentation in two tanks from two producers (B: industrial and P: hand-crafted). The prophage investigation was performed after a shotgun sequencing of whole genomic DNA, at each sampling time, through the exploration of prophage sequences in the bacterial metagenomic data. The number of reads was homogenous for all samples, with an amount of 2 × 107 reads per sample, except for C1B3 which had more reads than the others (8 × 107). The microbial composition of tanks B and P was of course accessible through these data, with tank P containing more reads assigned to Eukaryota than tank B (Figures S2 and S3, Table S2). Tank B presented a higher abundance in bacteria than tank P (Figure S2, Table S2). Within the bacteria, both tanks were dominated by the Proteobacteria phylum during the first three sampling times, while LAB appeared on the last sampling time (C1B4 and C2P4) (Figure S4).
After the phage prediction in the bacterial sequences with VIBRANT, 1603 contigs of putative phages were identified (114 were initially predicted as temperate and 1489 as lytic), the vast majority with a low- or medium-quality level. Among these contigs, only two complete prophages were identified, NODE_55 and NODE_129. The prophage NODE_55 was of 46,931 bases in length and matched with a Podoviridae CT3IJ1 (identity of 93%, coverage <1%, accession number BK023809.1) after a BLAST search against the NCBI virus database. The prophage NODE_129 was of 37,572 bases in length and matched with a Siphoviridae CTEVG3 (identity of 81%, coverage 16%, accession number BK027610.1) after a BLAST against the NCBI virus database. The 1603 phage contigs identified with VIBRANT were subjected to BLAST against the NCBI non-redundant nucleotide database (E-value < 10−16 and percentage of identity >90%) to generate information about the potential host. A total of 429 contigs (417 initially identified as lytic and 12 as temperate) gave no host identification according to the selected criteria, and were eliminated from the analysis of the prophages. So, 1174 contigs were retained, among them, 101 classified as temperate and 1073 as lytic by VIBRANT. Further analysis finally showed that all phage contigs matched with bacterial genomes and, therefore, were confirmed prophages. These results showed that the analysis of the life cycle by VIBRANT was a bit more uncertain in this case, probably due to the low-quality level of identification obtained.
A heatmap representation was used to visualise the dynamics of the temperate phage contigs inside bacterial metagenomes during the first month of the fermentation process for producer B (Figure 1). No heatmap could be made for producer P because the abundance was too small (RPKM max = 11.4 and 1593 values under an RPKM of 5.0 almost all equal to 0). Overall, the abundance of prophages in producer B samples tended to increase for some hosts, while other prophages fluctuated or decreased. For example, the occurrence of contig NODE_10382, affiliated to the Rahnella aqualitis species, increased overtime (RPKM of 13.6, 39.5, 36.4 and 35.6 in C1B1, C1B2, C1B3 and C1B4, respectively) (Table S3). Contig NODE_10904, affiliated to Pantoea agglomerans, decreased after C1B1 (RPKM moving from 29.2 to 7.7, 9.0 and 9.7 in C1B1, C1B2, C1B3 and C1B4, respectively) and contig NODE_3329, associated to Kluyvera intermedia, varied over time (RPKM passing from 17.9 to 25.9, 20.1 and 17.1 in C1B1, C1B2, C1B3 and C1B4, respectively) (Table S3). As expected, these observations of prophage occurrence altogether followed the same patterns as the levels of abundance of the bacterial genera to which they were associated. For example, the genus Rahnella increased overtime (total abundance of 4%, 14%, 12% and 11% in C1B1, C1B2, C1B3 and C1B4, respectively), the genus Pantoea decreased overtime (total abundance of 15%, 4%, 5% and 5% in C1B1, C1B2, C1B3 and C1B4, respectively), and the genus Kluyvera varied overtime (total abundance of 7%, 11%, 10% and 8% in C1B1, C1B2, C1B3 and C1B4, respectively) (data not shown).
Furthermore, gene identification in all the phage contigs was carried out using PROKKA. It did not reveal any gene of potential interest, as more than 95% of the coding DNA sequences were identified as hypothetical proteins (data not shown).
In addition to eukaryotic and bacterial sequences, which included prophages, a marginal part of viral DNA was also recovered (only 0.08% of the total reads), indicating traces of viral particles in the tanks (Figure 2). Among this small part of the total sequences, 75% of the viral sequences in tank B belonged to phages of the Caudoviricetes class (former Caudovirales order) [33], 39% to the myophages (former Myoviridae family), 23 to 26% to the siphophages (former Siphoviridae family) and 6 to 8% to the podophages (former Podoviridae family) at all the time points (Figure 2). In tank P, phages of the Caudoviricetes class were also recovered (16 to 28% of myophages, 10 to 19% of siphophages and 1 to 7% of podophages) (Figure 2). In this tank, the most abundant recovered sequences were those of viruses belonging to the Mimiviridae family, infecting amoeba and protists (10 to 35%) (Figure 2).

4. Discussion

The interest in phageomes, has been growing in recent years [8]. However, only three studies have been strictly dedicated to the phageome of fermented foods to date [9,10,11], and one to the airborne virome in cheese-making factories [12]. No published study about the phageomes in fermented beverages is available so far. It has been shown that the phageome plays an essential role in the microbial structure and balance of ecosystems, this being especially described in the human gut microbiota [24]. In this context, it was interesting to investigate the phageome of a fermented beverage, cider, in order to assess the diversity and occurrence of free lytic phages in different cider samples throughout a fermentation process in an industrial producer (B) and in a hand-crafted producer (P). It appeared that extracting free phage DNA from cider is not easily achievable when adapting protocols used for other types of samples, such as faeces [2] and cheese [11]. Indeed, in these matrices, the level of phages is usually high (>109 particles per gram), and phage DNA extraction seems thus facilitated. When using these methods in cider samples spiked with exogenous phage UCMA 21115, no phage DNA was recovered when the phage levels were under 107 PFU/mL. To improve phage DNA extraction in cider, PVPP at 1% and 2% was used to limit the potential negative impact of polyphenols in cider, known to be present at high concentration levels [22]. This molecule was used in buffers for plant DNA extraction, also known for their high contents of phenolic compounds, and its use showed an improvement in DNA extraction [25]. No improvement in phage DNA extraction was achieved using PVPP in cider. Polyphenols can be a way to explain the low quantity of phages in cider due to their antiviral properties [34]. They can act either by affecting the host killing the activity of phages, such as for two phages (OE33PA and Vinitor162) infecting Oenococcus oeni in wine, either by interacting with the phage tail or competing with the host site of recognition [35].
Tangential flow filtration [36] and Fe-based flocculation [24], two classical methods used for the concentration of viruses from marine waters, were then tested to increase the likelihood of getting higher quantities of phages and thus of extracting phage DNA from larger volumes of cider samples. Even so, free phage DNA extraction could not be improved in cider. Considering these results, it appears that cider is a fermented matrix with probable low levels of free lytic phages, that are difficult to concentrate for extracting phageome DNA. This was corroborated in the current study as no lytic phage was isolated against 160 strains by a double layer agar plate, even after TFF and flocculation. It is probably the case for other fermented beverages, as very few lytic phages from fermented beverages are described in the literature to date. Two lytic phages targeting O. oeni (OE33PA and Vinitor162) and one lytic phage targeting Gluconobacter cerinus (GC1) have been recently described in wine [18,37,38]. Moreover, in a previous work, 120 samples (cider, apple must, crushed apples) were screened for phages targeting 160 potential strains from genera found in cider, and only one lytic siphophage UCMA 21115 was isolated [23]. This result strengthens the hypothesis that cider is a poor reservoir of free lytic phages.
Cider, due to its harsh environment, presents a less complex bacterial diversity than other fermented foods or habitats [22]. The low microbial diversity could favour the temperate lifestyle of phages (lysogeny of prophages) because it results in an absence of predator/prey dynamics [15]. As the phageome of cider seemed scarce, a particular attention was thus paid to the prophages in cider throughout the fermentation process of producer (B) and producer (P). To date, no study has been focused on the prophages in their entirety in fermented foods and beverages, even if this topic is becoming an emerging field of investigations. A recent study on wine samples from three wineries investigated the prevalence of prophages in O. oeni strains through the fermentation process [39]. The majority of studies investigated the distribution of prophages in bacterial genomes, as is the case for the Lactobacillaceae strains, members of the former Lactobacillus genus [40], and for the O. oeni strains [41]. These studies highlight that prophage diversity varies from one species to another, and that prophages are retrieved in many strains. Prophages are also recognised as markers of diversity within a species in the case of O. oeni, and as sources of horizontal gene transfer in Lactobacillus. Prophages are also known as bacterial fitness enhancers. For example, Streptococcus pyogenes prophage, carrying genes encoding exotoxins, increased colonisation during scarlet fever [42], and an Escherichia coli strain carrying a prophage with a gene encoding antibiotic resistance was more adapted to its environment under antibiotic pressure [43]. These examples demonstrate the importance of prophages on their host and the necessity to evaluate their impact and diversity in fermented foods and beverages, which is little documented to date at the scale of the whole bacterial community.
In the current study, a metagenomic approach was carried out to observe prophage signatures in bacterial metagenomes throughout the cider fermentation process. A total of 1174 putative prophage contigs were identified in silico in the bacterial metagenomes using VIBRANT software, such as has been done in cheese [44] and kefir [45]. Among them, two complete prophages were found, NODE_55 and NODE_129, matching with phages isolated from human gut metagenomic data. The majority of the prophages identified in the current study were related to bacterial genera classically associated with the apple microbiota, such as, for example, Citrobacter, Klebsiella, Pantoa, Pseudomonas and Xanthomonas [46,47]. The prophage dynamics in cider were correlated to their bacterial host dynamics over time. For example, some prophages, such as NODE_10382, increased, others, such as NODE_10904, decreased and some others, such as NODE_3329, fluctuated during the fermentation process. These observations were correlated to the level of bacteria of the same genera with which they were associated at each fermentation time point. The increase in the frequency of prophages over time could tend to show that lysogeny had probably been established and was predominant. Conversely, a decrease in prophage frequency could be associated to a decrease in lysogeny. Indeed, this decrease could be due to the fact that the prophages have entered a lytic cycle, and consequently, are less present in their hosts [48,49,50]. In the current study, it was not possible to determine if these prophages were active or not with this method of analysis and with the encountered difficulties to extract free lytic viral particles, as described above. In a study dealing with the virome of pigs, active and inactive prophages were distinguished with bioinformatic tools after the extraction of lytic viral particles [51]. Another possible method could be to induce prophages with mitomycine C as it has been performed on gut microbiota to study active prophages [52], bearing in mind a bias due to the fact that not all prophages are inducible. The main barrier for these types of approaches in cider, and probably in other fermented beverages, is the problematic extraction of phage DNA, due to their very low occurrence, and also to the particular physicochemical conditions encountered in these fermented beverages. Positive and negative correlations could be made in future experiments between lytic phages, prophages and bacterial proportions in cider throughout the fermentation process by combining the analyses of metabolites and food properties, as it has been described in cheese, for example [44]. For a better view of these dynamics in cider, and especially with LAB, it would be interesting to extend the timespan of the study looking at longer fermentation processes, such as cider intended to be used for the making of spirits and vinegars [53].

5. Conclusions

This study revealed the low occurrence of free lytic phages, and thus of phageome, in cider. This fermented beverage appears to be a poor reservoir of lytic phages in contrast to other food ecosystems, such as cheeses [11]. The presence of phages in cider is more likely to occur essentially as prophages throughout the fermentation process. Besides the fact that cider does not contain many phages, a number of questions are raised about fermented beverages. What is special about this type of ecosystem compared to other hostile environments where phages are found? Why have lytic phages not been able, at first sight, to adapt to this environment, even though it is quite rich in bacteria? Are polyphenols implicated in this situation? All these questions remain unanswered for the time being, and show the importance of looking at them in more details in order to better understand the place of phages in these ecosystems.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/microorganisms10061203/s1, Figure S1: Agarose gels of phage DNA extraction from cider adjusted to pH 5.5 spiked with phage UCMA 21115. Figure S2: Microbial taxonomic assignations at the domain level during the cider fermentation process for two producers (B: industrial and P: hand-crafted). Figure S3: Eukaryota taxonomic assignations at the genus level during the cider fermentation process for two producers (B: industrial and P: hand-crafted). Figure S4: Bacterial taxonomic assignations at the family level during cider fermentation process for two producers (B: industrial and P: hand-crafted). Table S1: Percentage of the number of reads attributed to Super-kingdoms in NCBI. Table S2: Bacterial strains used to isolate bacteriophages. Table S3: Occurrence of prophages for producer B and P.

Author Contributions

Conceptualisation: P.L., M.D. and N.D. Methodology: P.L., M.D. and N.D. Formal analysis: O.R., C.M. and P.L. Data curation: O.R., C.M., P.L., M.D. and N.D. Writing—original draft preparation: P.L., M.D. and N.D. Writing—review and editing: P.L., M.D., O.R., C.M. and N.D. Supervision: M.D. and N.D. Project administration: M.D and N.D. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported by the French Ministère de l’Enseignement Supérieur, de la Recherche et de l’Innovation under a PhD Grant MENRT.

Data Availability Statement

Publicly available datasets were analysed in this study. This data can be found in the NCBI SRA repository, reference number PRJNA803977.

Acknowledgments

The authors would like to thank Céline Caillard and Erwan Renard for their technical support. We are grateful to the INRAE MIGALE bioinformatics facility (MIGALE, INRAE, 2020. Migale bioinformatics Facility, doi:10.15454/1.5572390655343293E12) for providing computing and storage resources.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Manrique, P.; Bolduc, B.; Walk, S.T.; van der Oost, J.; de Vos, W.M.; Young, M.J. Healthy Human Gut Phageome. Proc. Natl. Acad. Sci. USA 2016, 113, 10400–10405. [Google Scholar] [CrossRef] [Green Version]
  2. Shkoporov, A.N.; Ryan, F.J.; Draper, L.A.; Forde, A.; Stockdale, S.R.; Daly, K.M.; McDonnell, S.A.; Nolan, J.A.; Sutton, T.D.S.; Dalmasso, M.; et al. Reproducible Protocols for Metagenomic Analysis of Human Faecal Phageomes. Mi-Crobiome 2018, 6, 68. [Google Scholar] [CrossRef] [PubMed]
  3. Dalmasso, M.; Hill, C.; Ross, R.P. Exploiting Gut Bacteriophages for Human Health. Trends Microbiol. 2014, 22, 399–405. [Google Scholar] [CrossRef] [PubMed]
  4. Carr, V.R.; Shkoporov, A.; Gomez-Cabrero, D.; Mullany, P.; Hill, C.; Moyes, D.L. The Human Oral Phageome Is Highly Diverse and Rich in Jumbo Phages. bioRxiv 2020. [Google Scholar] [CrossRef]
  5. Ma, Y.; Marais, A.; Lefebvre, M.; Theil, S.; Svanella-Dumas, L.; Faure, C.; Candresse, T. Phytovirome Analysis of Wild Plant Populations: Comparison of Double-Stranded RNA and Virion-Associated Nucleic Acid Metagenomic Approaches. J. Virol. 2019, 94, e01462-19. [Google Scholar] [CrossRef]
  6. Roux, S.; Brum, J.R.; Dutilh, B.E.; Sunagawa, S.; Duhaime, M.B.; Loy, A.; Poulos, B.T.; Solonenko, N.; Lara, E.; Poulain, J.; et al. Ecogenomics and Potential Biogeochemical Impacts of Globally Abundant Ocean Viruses. Nature 2016, 537, 689–693. [Google Scholar] [CrossRef] [Green Version]
  7. Zhong, Z.-P.; Tian, F.; Roux, S.; Gazitúa, M.C.; Solonenko, N.E.; Li, Y.-F.; Davis, M.E.; Van Etten, J.L.; Mosley-Thompson, E.; Rich, V.I.; et al. Glacier Ice Archives Nearly 15,000-Year-Old Microbes and Phages. Microbiome 2021, 9, 160. [Google Scholar] [CrossRef]
  8. Ledormand, P.; Desmasures, N.; Dalmasso, M. Phage Community Involvement in Fermented Beverages: An Open Door to Technological Advances? Crit. Rev. Food Sci. Nutr. 2020, 61, 2911–2920. [Google Scholar] [CrossRef]
  9. Jung, M.-J.; Kim, M.-S.; Yun, J.-H.; Lee, J.-Y.; Kim, P.S.; Lee, H.-W.; Ha, J.-H.; Roh, S.W.; Bae, J.-W. Viral Community Predicts the Geographical Origin of Fermented Vegetable Foods More Precisely than Bacterial Community. Food Microbiol. 2018, 76, 319–327. [Google Scholar] [CrossRef]
  10. Park, E.-J.; Kim, K.-H.; Abell, G.C.; Kim, M.-S.; Roh, S.W.; Bae, J.-W. Metagenomic Analysis of the Viral Communities in Fermented Foods. Appl. Environ. Microbiol. 2011, 77, 1284–1291. [Google Scholar] [CrossRef] [Green Version]
  11. Dugat-Bony, E.; Lossouarn, J.; De Paepe, M.; Sarthou, A.-S.; Fedala, Y.; Petit, M.-A.; Chaillou, S. Viral Metagenomic Analysis of the Cheese Surface: A Comparative Study of Rapid Procedures for Extracting Viral Particles. Food Microbiol. 2020, 85, 103278. [Google Scholar] [CrossRef] [PubMed]
  12. Colombo, S.; Arioli, S.; Gargari, G.; Neri, E.; Della Scala, G.; Mora, D. Characterization of Airborne Viromes in Cheese Production Plants. J. Appl. Microbiol. 2018, 125, 1444–1454. [Google Scholar] [CrossRef] [PubMed]
  13. Feiner, R.; Argov, T.; Rabinovich, L.; Sigal, N.; Borovok, I.; Herskovits, A.A. A New Perspective on Lysogeny: Prophages as Active Regulatory Switches of Bacteria. Nat. Rev. Microbiol. 2015, 13, 641–650. [Google Scholar] [CrossRef] [PubMed]
  14. Howard-Varona, C.; Lindback, M.M.; Bastien, G.E.; Solonenko, N.; Zayed, A.A.; Jang, H.; Andreopoulos, B.; Brewer, H.M.; del Rio, T.G.; Adkins, J.N.; et al. Phage-Specific Metabolic Reprogramming of Virocells. ISME J. 2020, 14, 881–895. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  15. Mills, S.; Shanahan, F.; Stanton, C.; Hill, C.; Coffey, A.; Ross, R.P. Movers and Shakers: Influence of Bacteriophages in Shaping the Mammalian Gut Microbiota. Gut Microbes 2013, 4, 4–16. [Google Scholar] [CrossRef] [PubMed]
  16. LeLièvre, V.; Besnard, A.; Schlusselhuber, M.; Desmasures, N.; Dalmasso, M. Phages for Biocontrol in Foods: What Opportunities for Salmonella sp. Control along the Dairy Food Chain? Food Microbiol. 2019, 78, 89–98. [Google Scholar] [CrossRef]
  17. Pietrysiak, E.; Smith, S.; Ganjyal, G.M. Food Safety Interventions to Control Listeria monocytogenes in the Fresh Apple Packing Industry: A Review. Compr. Rev. Food Sci. Food Saf. 2019, 18, 1705–1726. [Google Scholar] [CrossRef] [Green Version]
  18. Philippe, C.; Krupovic, M.; Jaomanjaka, F.; Claisse, O.; Petrel, M.; Le Marrec, C. Bacteriophage GC1, a Novel Tectivirus Infecting Gluconobacter cerinus, an Acetic Acid Bacterium Associated with Wine-Making. Viruses 2018, 10, 39. [Google Scholar] [CrossRef] [Green Version]
  19. Deasy, T.; Mahony, J.; Neve, H.; Heller, K.J.; Van Sinderen, D. Isolation of a Virulent Lactobacillus brevis Phage and Its Application in the Control of Beer Spoilage. J. Food Prot. 2011, 74, 2157–2161. [Google Scholar] [CrossRef]
  20. Mahony, J.; Van Sinderen, D. Current Taxonomy of Phages Infecting Lactic Acid Bacteria. Front. Microbiol. 2014, 5, 7. [Google Scholar] [CrossRef] [Green Version]
  21. Misery, B.; Legendre, P.; Rue, O.; Bouchart, V.; Guichard, H.; Laplace, J.M.; Cretenet, M. Diversity and Dynamics of Bacterial and Fungal Communities in Cider for Distillation. Int. J. Food Microbiol. 2021, 339, 108987. [Google Scholar] [CrossRef]
  22. Cousin, F.J.; Le Guellec, R.; Schlusselhuber, M.; Dalmasso, M.; Laplace, J.-M.; Cretenet, M. Microorganisms in Fermented Apple Beverages: Current Knowledge and Future Directions. Microorganisms 2017, 5, 39. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  23. Ledormand, P.; Desmasures, N.; Bernay, B.; Goux, D.; Monnet, C.; Dalmasso, M. Molecular Approaches to Uncover Phage-Lactic Acid Bacteria Interactions in a Model Community Simulating Fermented Beverages. Food Microbiol. 2022, in press. [CrossRef]
  24. Draper, L.A.; Ryan, F.J.; Dalmasso, M.; Casey, P.G.; McCann, A.; Velayudhan, V.; Ross, R.P.; Hill, C. Autochthonous Faecal Virome Transplantation (FVT) Reshapes the Murine Microbiome after Antibiotic Perturbation. bioRxiv 2019. [Google Scholar] [CrossRef] [Green Version]
  25. Kasajima, I.; Sasaki, K.; Tanaka, Y.; Terakawa, T.; Ohtsubo, N. Large-Scale Extraction of Pure DNA from Mature Leaves of Cyclamen Persicum Mill. and Other Recalcitrant Plants with Alkaline Polyvinylpolypyrrolidone (PVPP). Sci. Hortic. 2013, 164, 65–72. [Google Scholar] [CrossRef]
  26. John, S.G.; Mendez, C.B.; Deng, L.; Poulos, B.; Kauffman, A.K.M.; Kern, S.; Brum, J.; Polz, M.F.; Boyle, E.A.; Sullivan, M.B. A Simple and Efficient Method for Concentration of Ocean Viruses by Chemical Flocculation. Environ. Microbiol. Rep. 2011, 3, 195–202. [Google Scholar] [CrossRef] [Green Version]
  27. Andrews, S. Fastqc: A Quality Control Tool for High Throughput Sequence Data; Babraham Bioinformatics. 2010. Available online: https://www.bioinformatics.babraham.ac.uk/projects/fastqc/ (accessed on 8 June 2022).
  28. Chen, S.; Zhou, Y.; Chen, Y.; Gu, J. Fastp: An Ultra-Fast All-in-One FASTQ Preprocessor. Bioinformatics 2018, 34, i884–i890. [Google Scholar] [CrossRef]
  29. Bankevich, A.; Nurk, S.; Antipov, D.; Gurevich, A.A.; Dvorkin, M.; Kulikov, A.S.; Lesin, V.M.; Nikolenko, S.I.; Pham, S.; Prjibelski, A.D.; et al. SPAdes: A New Genome Assembly Algorithm and Its Applications to Single-Cell Sequencing. J. Comput. Biol. 2012, 19, 455–477. [Google Scholar] [CrossRef] [Green Version]
  30. Menzel, P.; Ng, K.L.; Krogh, A. Fast and Sensitive Taxonomic Classification for Metagenomics with Kaiju. Nat. Commun. 2016, 7, 11257. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  31. Kieft, K.; Zhou, Z.; Anantharaman, K. VIBRANT: Automated Recovery, Annotation and Curation of Microbial Viruses, and Evaluation of Viral Community Function from Genomic Sequences. Microbiome 2020, 8, 90. [Google Scholar] [CrossRef]
  32. Li, W.; Godzik, A. Cd-Hit: A Fast Program for Clustering and Comparing Large Sets of Protein or Nucleotide Sequences. Bioinformatics 2006, 22, 1658–1659. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Turner, D.; Kropinski, A.M.; Adriaenssens, E.M. A Roadmap for Genome-Based Phage Taxonomy. Viruses 2021, 13, 506. [Google Scholar] [CrossRef] [PubMed]
  34. Catel-Ferreira, M.; Tnani, H.; Hellio, C.; Cosette, P.; Lebrun, L. Antiviral Effects of Polyphenols: Development of Bio-Based Cleaning Wipes and Filters. J. Virol. Methods 2015, 212, 1–7. [Google Scholar] [CrossRef] [PubMed]
  35. Philippe, C.; Chaïb, A.; Jaomanjaka, F.; Cluzet, S.; Lagarde, A.; Ballestra, P.; Decendit, A.; Petrel, M.; Claisse, O.; Goulet, A.; et al. Wine Phenolic Compounds Differently Affect the Host-Killing Activity of Two Lytic Bacteriophages Infecting the Lactic Acid Bacterium Oenococcus oeni. Viruses 2020, 12, 1316. [Google Scholar] [CrossRef] [PubMed]
  36. Cai, L.; Yang, Y.; Jiao, N.; Zhang, R. Evaluation of Tangential Flow Filtration for the Concentration and Separation of Bacteria and Viruses in Contrasting Marine Environments. PLoS ONE 2015, 10, e0136741. [Google Scholar] [CrossRef]
  37. Jaomanjaka, F.; Claisse, O.; Blanche-Barbat, M.; Petrel, M.; Ballestra, P.; Le Marrec, C. Characterization of a New Virulent Phage Infecting the Lactic Acid Bacterium Oenococcus oeni. Food Microbiol. 2016, 54, 167–177. [Google Scholar] [CrossRef]
  38. Philippe, C.; Chaïb, A.; Jaomanjaka, F.; Claisse, O.; Lucas, P.M.; Samot, J.; Cambillau, C.; Le Marrec, C. Characterization of the First Virulent Phage Infecting Oenococcus oeni, the Queen of the Cellars. Front. Microbiol. 2021, 11, 596541. [Google Scholar] [CrossRef]
  39. Chaïb, A.; Claisse, O.; Delbarre, E.; Bosviel, J.; Le Marrec, C. Assessment of the Lysogenic Status in the Lactic Acid Bacterium O. oeni during the Spontaneous Malolactic Fermentation of Red Wines. Food Microbiol. 2022, 103, 103947. [Google Scholar] [CrossRef] [PubMed]
  40. Pei, Z.; Sadiq, F.A.; Han, X.; Zhao, J.; Zhang, H.; Ross, R.P.; Lu, W.; Chen, W. Comprehensive Scanning of Prophages in Lactobacillus: Distribution, Diversity, Antibiotic Resistance Genes, and Linkages with CRISPR-Cas Systems. mSystems 2021, 6, e0121120. [Google Scholar] [CrossRef]
  41. Claisse, O.; Chaïb, A.; Jaomanjaka, F.; Philippe, C.; Barchi, Y.; Lucas, P.M.; Le Marrec, C. Distribution of Prophages in the Oenococcus oeni Species. Microorganisms 2021, 9, 856. [Google Scholar] [CrossRef]
  42. Brouwer, S.; Barnett, T.C.; Ly, D.; Kasper, K.J.; De Oliveira, D.M.P.; Rivera-Hernandez, T.; Cork, A.J.; McIntyre, L.; Jespersen, M.G.; Richter, J.; et al. Prophage Exotoxins Enhance Colonization Fitness in Epidemic Scarlet Fever-Causing Streptococcus pyogenes. Nat. Commun. 2020, 11, 5018. [Google Scholar] [CrossRef] [PubMed]
  43. Wendling, C.C.; Refardt, D.; Hall, A.R. Fitness Benefits to Bacteria of Carrying Prophages and Prophage-Encoded Antibiotic-Resistance Genes Peak in Different Environments. Evolution 2021, 75, 515–528. [Google Scholar] [CrossRef] [PubMed]
  44. Yang, C.; You, L.; Kwok, L.-Y.; Jin, H.; Peng, J.; Zhao, Z.; Sun, Z. Strain-Level Multiomics Analysis Reveals Significant Variation in Cheeses from Different Regions. LWT 2021, 151, 112043. [Google Scholar] [CrossRef]
  45. Tenorio-Salgado, S.; Castelán-Sánchez, H.G.; Dávila-Ramos, S.; Huerta-Saquero, A.; Rodríguez-Morales, S.; Merino-Pérez, E.; Roa de la Fuente, L.F.; Solis-Pereira, S.E.; Pérez-Rueda, E.; Lizama-Uc, G. Metagenomic Analysis and Antimicrobial Activity of Two Fermented Milk Kefir Samples. Microbiol. Open 2021, 10, e1183. [Google Scholar] [CrossRef] [PubMed]
  46. Abadias, M.; Cañamás, T.P.; Asensio, A.; Anguera, M.; Viñas, I. Microbial Quality of Commercial “Golden Delicious” Apples throughout Production and Shelf-Life in Lleida (Catalonia, Spain). Int. J. Food Microbiol. 2006, 108, 404–409. [Google Scholar] [CrossRef]
  47. Liu, J.; Abdelfattah, A.; Norelli, J.; Burchard, E.; Schena, L.; Droby, S.; Wisniewski, M. Apple Endophytic Microbiota of Different Rootstock/Scion Combinations Suggests a Genotype-Specific Influence. Microbiome 2018, 6, 18. [Google Scholar] [CrossRef] [PubMed]
  48. Kim, M.-S.; Bae, J.-W. Lysogeny Is Prevalent and Widely Distributed in the Murine Gut Microbiota. ISME J. 2018, 12, 1127–1141. [Google Scholar] [CrossRef] [Green Version]
  49. Chevallereau, A.; Pons, B.J.; van Houte, S.; Westra, E.R. Interactions between Bacterial and Phage Communities in Natural Environments. Nat. Rev. Microbiol. 2021, 20, 49–62. [Google Scholar] [CrossRef]
  50. Howard-Varona, C.; Hargreaves, K.R.; Abedon, S.T.; Sullivan, M.B. Lysogeny in Nature: Mechanisms, Impact and Ecology of Temperate Phages. ISME J. 2017, 11, 1511–1520. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  51. Billaud, M.; Lamy-Besnier, Q.; Lossouarn, J.; Moncaut, E.; Dion, M.B.; Moineau, S.; Traoré, F.; Le Chatelier, E.; Denis, C.; Estelle, J.; et al. Analysis of Viromes and Microbiomes from Pig Fecal Samples Reveals That Phages and Prophages Rarely Carry Antibiotic Resistance Genes. ISME Commun. 2021, 1, 55. [Google Scholar] [CrossRef]
  52. Zünd, M.; Ruscheweyh, H.-J.; Field, C.M.; Meyer, N.; Cuenca, M.; Hoces, D.; Hardt, W.-D.; Sunagawa, S. High Throughput Sequencing Provides Exact Genomic Locations of Inducible Prophages and Accurate Phage-to-Host Ratios in Gut Microbial Strains. Microbiome 2021, 9, 77. [Google Scholar] [CrossRef] [PubMed]
  53. Trček, J.; Mahnič, A.; Rupnik, M. Diversity of the Microbiota Involved in Wine and Organic Apple Cider Submerged Vinegar Production as Revealed by DHPLC Analysis and Next-Generation Sequencing. Int. J. Food Microbiol. 2016, 223, 57–62. [Google Scholar] [CrossRef] [PubMed]
Microorganisms 10 01203 g001 550
Figure 1. Heatmap showing the dynamics of the most abundant prophages retrieved during the first month of the fermentation process for producer B. C1 indicates the tank ID, and B1, B2, B3 and B4 are the sampling points during the fermentation process, in reference to Table 1. NODE referred to the identified prophage and the mentioned bacterial identifications represent the best host species of the prophage predicted after blast (E-value < 10−16; % of identity >85%). RPKM value above 5.0 was considered for the construction of the heatmap. Green to red: lowest to highest abundance.
Figure 1. Heatmap showing the dynamics of the most abundant prophages retrieved during the first month of the fermentation process for producer B. C1 indicates the tank ID, and B1, B2, B3 and B4 are the sampling points during the fermentation process, in reference to Table 1. NODE referred to the identified prophage and the mentioned bacterial identifications represent the best host species of the prophage predicted after blast (E-value < 10−16; % of identity >85%). RPKM value above 5.0 was considered for the construction of the heatmap. Green to red: lowest to highest abundance.
Microorganisms 10 01203 g001
Microorganisms 10 01203 g002 550
Figure 2. Virus taxonomic assignations at the family level during cider fermentation process for two producers (B: industrial and P: hand-crafted). C1 and C2 indicate the tank ID, and B1, B2, B3, B4, P1, P2, P3 and P4 are the sampling points during the fermentation process, in reference to Table 1.
Figure 2. Virus taxonomic assignations at the family level during cider fermentation process for two producers (B: industrial and P: hand-crafted). C1 and C2 indicate the tank ID, and B1, B2, B3, B4, P1, P2, P3 and P4 are the sampling points during the fermentation process, in reference to Table 1.
Microorganisms 10 01203 g002
Table
Table 1. Cider samples collected for the study of the phageome and for the prophage investigation.
Table 1. Cider samples collected for the study of the phageome and for the prophage investigation.
Sample IDProducer ID (French Department)Fermentation Time (Days)Tank ID—Tank Volume (hL)Sugar Concentration kg/m3pH
C1B1B (Calvados)0C1—12010604.02
C1B2B (Calvados)7C1—12010564.05
C1B3B (Calvados)14C1—12010484.03
C1B4B (Calvados)27C1—12010364.01
C2P1P (Manche)0C2—1010563.32
C2P2P (Manche)12C2—1010493.35
C2P3P (Manche)25C2—1010403.52
C2P4P (Manche)36C2—1010363.58
Table
Table 2. Overview of the conditions tested for the collection of the phageome from cider. Roman numerals from I to IX indicate methodological steps referred to in the text.
Table 2. Overview of the conditions tested for the collection of the phageome from cider. Roman numerals from I to IX indicate methodological steps referred to in the text.
Tested Conditions to Collect Viral Particles from Cider
IIIIIIIVVVIVIIVIII
Tangential Flow Filtration		IX
Flocculation with FeCl3
Tested volumes of cider (mL)5020, 50, 1000400
Sample spiked with phage UCMA 21115 (PFU/mL)Yes (104–109) Yes (105)Yes (104–108)
pH adjusted to 5.5 Yes Yes
Precipitation with 10% PEG-8000 and 0.5 M NaCl YesYes YesYesYes
Centrifugation stepYesYesYesYesYesYesYes
DNA extraction fromPelletSupernatantPelletPelletSupernatantPelletPelletRetentateFlocculate
Table
Table 3. Efficiency of iron-based flocculation to recover phage UCMA 21115 from cider samples.
Table 3. Efficiency of iron-based flocculation to recover phage UCMA 21115 from cider samples.
Inoculated Quantity of Phage UCMA 21115 in 400 mL of Cider (i.e., Concentration)Retrieved Quantity of Phage UCMA 21115 after Flocculation in Cider
4 × 1010
(1 × 108 PFU/mL)		8.5 × 108
4 × 109
(1 × 107 PFU/mL)		9.9 × 107
4 × 108
(1 × 106 PFU/mL)		6.6 × 106
4 × 107
(1 × 105 PFU/mL)		7.6 × 105
4 × 106
(1 × 104 PFU/mL)		5.0 × 103
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Ledormand, P.; Desmasures, N.; Midoux, C.; Rué, O.; Dalmasso, M. Investigation of the Phageome and Prophages in French Cider, a Fermented Beverage. Microorganisms 2022, 10, 1203. https://doi.org/10.3390/microorganisms10061203

AMA Style

Ledormand P, Desmasures N, Midoux C, Rué O, Dalmasso M. Investigation of the Phageome and Prophages in French Cider, a Fermented Beverage. Microorganisms. 2022; 10(6):1203. https://doi.org/10.3390/microorganisms10061203

Chicago/Turabian Style

Ledormand, Pierre, Nathalie Desmasures, Cédric Midoux, Olivier Rué, and Marion Dalmasso. 2022. "Investigation of the Phageome and Prophages in French Cider, a Fermented Beverage" Microorganisms 10, no. 6: 1203. https://doi.org/10.3390/microorganisms10061203

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