Targeted Metagenomics of Retting in Flax: The Beginning of the Quest to Harness the Secret Powers of the Microbiota
1Univ. Lille, CNRS, UMR 8576 – UGSF – Unité de Glycobiologie Structurale et Fonctionnelle, Lille, France
2Université de Reims Champagne Ardenne, INRAE, UMR FARE A 614, Reims, France
*Correspondence: Sébastien Grec, sebastien.grec@univ-lille.frAbstract
The mechanical and chemical properties of natural plant fibers are determined by many different factors, both intrinsic and extrinsic to the plant, during growth but also after harvest. A better understanding of how all these factors exert their effect and how they interact is necessary to be able to optimize fiber quality for use in different industries. One important factor is the post-harvest process known as retting, representing the first step in the extraction of bast fibers from the stem of species such as flax and hemp. During this process microorganisms colonize the stem and produce hydrolytic enzymes that target cell wall polymers thereby facilitating the progressive destruction of the stem and fiber bundles. Recent advances in sequencing technology have allowed researchers to implement targeted metagenomics leading to a much better characterization of the microbial communities involved in retting, as well as an improved understanding of microbial dynamics. In this paper we review how our current knowledge of the microbiology of retting has been improved by targeted metagenomics and discuss how related ‘-omics’ approaches might be used to fully characterize the functional capability of the retting microbiome.
Introduction
Natural fibers from different plant species have long been used by man to make textiles and are now being increasingly exploited as a viable replacement for synthetic fibers in composite materials. From a biological point of view the fibers used in textiles and composites are single cells, much longer than they are wide and characterized by the presence of a thick secondary cell wall. Such structures can be found in the xylem (wood fibers) and/or associated with the phloem in the outer tissues of the stem of non-woody plants (e.g., the bast fibers of flax, hemp, ramie), as well as in the walls of the fruits or the leaves of some species (e.g., cotton, kapok, sisal). In bast fiber species, the individual fiber cells (elementary fibers) are grouped together to form so-called fiber bundles.
The ‘quality’ of the fabricated textiles and natural fiber composites (NFCs) is determined by both the mechanical and chemical properties of the isolated fibers/fiber bundles and the subsequent industrial transformation process (Müssig, 2010; Summerscales et al., 2010; Bourmaud et al., 2018), and researchers have therefore explored the different factors influencing the quality of the finished products.
It is generally accepted that the mechanical and chemical properties of plant fibers result from fiber morphology (e.g., length, diameter), cell wall composition (i.e., what polymers are present and in what quantities) and organization (i.e., where in the cell wall the polymers are located and how they are organized and interact with one another). At the biological level, these aspects are determined by the combined and coordinated spatio-temporal expression of several hundreds of genes, and a major challenge is therefore to identify which ones are involved in this process and to understand their role(s). As a result, genomic and genetic studies aimed at improving fiber plants have so far mainly focused on genome sequencing and the identification of genes/traits associated with various agronomic traits (e.g., plant height, fiber yield, fiber diameter) or cell wall construction (e.g., cellulose/pectin/lignin biosynthesis). Flax was the second fiber plant to have its genome sequenced (Wang et al., 2012) after hemp (van Bakel et al., 2011) and can be accessed on the phytozome public database1. The sequences of other fiber plants followed a few years later, most likely as a result of the size and/or complexity of the genome with cotton, jute and ramie being sequenced, respectively in 2015 (Li et al., 2015), 2016 (Yuan et al., 2016), 2017 (Sarkar et al., 2017), and 2018 (Luan et al., 2018). In parallel, a number of whole genome transcriptomics studies by microarrays and more recently RNAseq, together with targeted qRT-PCR and in situ hybridization, have enabled the identification of genes likely to be involved in cell wall polymer biosynthesis in flax (Roach and Deyholos, 2007; Fenart et al., 2010; Huis et al., 2012; Chantreau et al., 2015; Zhang and Deyholos, 2016; Gorshkov et al., 2017; le Roy et al., 2017). Despite the identification of numerous candidates, examples of functional validation of cell wall genes in flax, as in other fiber species – are relatively limited despite the fact that this species can be genetically transformed and mutant populations are available (Wróbel-Kwiatkowska et al., 2007; Day et al., 2009; Chantreau et al., 2013, 2014).
Linking fiber phenotype (morphology, composition and organization) to gene expression profiles in field-grown flax is an extremely challenging task as these profiles are continually modified during plant growth according to developmental and environmental cues (Figure 1). Both microarrays and RNAseq have been used to investigate modifications in gene expression in response to drought stress (Dash et al., 2014), saline and alkaline stress (Yu et al., 2014), and infection by the fungus Fusarium oxysporum (Galindo-González and Deyholos, 2016). In both cases, comparison of the transcriptomes from control plants and stressed plants revealed significant changes in the expression of a number of genes likely to have a direct/indirect impact on fiber phenotype.
The situation is further complicated by the fact that flax fibers are extracted from the stem by mechanical defibering that may alter their mechanical and chemical properties especially at the level of so-called ‘technical fibers’, corresponding to whole or fragmented fiber bundles. Furthermore, mechanical extraction itself is generally preceded by a retting step that can also modify fiber properties if not managed properly (Md.Tahir et al., 2011; Liu et al., 2017).
In field-/dew-retting, plants are pulled (uprooted) and left on the ground for several weeks during which they are colonized by microorganisms (e.g., fungi and bacteria) that produce cell wall degrading enzymes contributing to the progressive de-solidification of fiber bundles from the other stem tissues and partial dissociation of the fiber bundles (Figure 2A). Fibers can also be de-solidified by water-retting, a process in which flax plants are placed into water tanks and colonized mainly by anaerobic bacteria (Tamburini et al., 2003; Zhao et al., 2016). In both cases these natural biological processes affect fiber homogeneity and mechanical performance.
Until recently, our knowledge of the different microorganisms present during retting, the dynamics of colonization, and the complexity of the cell wall degrading enzyme arsenal has been severely limited by our incapacity to cultivate and study the great majority of these species. Furthermore, we know hardly anything about how assembly, activities (i.e., cell-wall degrading capacity) of the retting microbial community and interactions among members are affected by environmental conditions or by plant genotype (Figure 1). The fact that such considerations are important is demonstrated by the results of several studies that have shown that both abiotic factors (e.g., climate, soil properties) and biotic factors (microorganisms) continue to affect the phenotype of plant fibers after harvest during the process of retting (Martin et al., 2013; Liu et al., 2015; Mazian et al., 2018; Bleuze et al., 2020; Chabbert et al., 2020). As a result, retting of fiber species is largely based upon the farmer’s experience and remains a biological black box preventing the development of more objective approaches.
Currently, rapid advances in high-throughput sequencing (HTS) technologies are allowing us to exploit the rapidly expanding field of metagenomics to improve our understanding of retting in flax and other fiber plants (Djemiel et al., 2017; Chabbert et al., 2020). In this paper we review how the application of this approach promises to make a major contribution to the understanding of this complex process and we discuss the perspectives of how related strategies such as metatranscriptomics and metaproteomics could also lead to further advances.
The Retting of Flax, a Key Step for the Obtention of Quality Fibers
Retting – Definition and Types
One of the first definitions of retting is as follows: ‘Retting refers to the operation that textile plants are subjected to in order to free the fibers of the liber from the gum-resinous substance which binds them and keeps them attached to the woody stem of the plant’ (Renouard, 1890). The ‘art’ of retting is all about compromise: under-retting will give coarse technical fibers, consisting of several associated individual fibers and contaminated by wood debris and external tissues (cortical parenchyma and cutinized epidermis; Goodman et al., 2002), while over-retting will reduce the resistance of the fibers by affecting their integral structure (Rosemberg and de França, 1967; Akin et al., 1998).
Generally, two main types of retting are used: “water retting” and “field-/dew-retting.” For water retting, flax stems are harvested, grouped in bundles and submerged in water. Historically, this was done in natural basins (e.g., lakes, rivers or dams) for 5–7 days, followed by a period where stems were dried directly on the ground for one to 2 weeks (Akin, 2010). More recently, artificial pits or tanks have been used to wash the stems with clean water to remove residues. This technique makes it possible to control parameters influencing bacterial development including aeration and temperature, and also allows the inoculation of selected bacterial strains. Water-retting depends mainly on the action of anaerobic bacteria that colonize the stem and is related to fermentation (Donaghy et al., 1990). This method generates high quality flax fibers but is also a source of environmental pollution (Md.Tahir et al., 2011). In field-/dew-retting, flax plants are mechanically up-rooted (pulled) by specialized harvesters and the stems placed directly on the field in piles to form swathes (Figure 1; Meijer et al., 1995). Subsequently, the humidity brought by the morning dew and the alternation of rain and heat, favors the development of microorganisms (mainly bacteria and fungi) initially present on the stems and the colonization by the microflora from the soil on which the swathes are placed (Sharma and Faughey, 1999; Akin, 2010; Djemiel et al., 2017). During retting, the intervention of the farmer involves turning the swathes halfway through the process in order to ensure uniform retting over the entire swathe height, and surveying the progression of the process (Franck, 2005; Akin, 2013).
More recently, various alternative methods using commercial enzyme cocktails mainly containing pectinases and xylanases have been investigated in an attempt to provide better control of the process and therefore improve fiber yield and quality (Henriksson et al., 1997; Zhang et al., 2000; Akin et al., 2001). However, their use remains limited mainly because of the high cost of producing these enzymes in bioreactors.
Retting and Fiber Cell Walls
Individual flax fiber cells (elementary fibers) are grouped together to form fiber bundles that are found in the stem outer tissues surrounding the xylem and pith that constitute the inner tissues (Figure 2A). The bundles are located beneath the epidermis and between the cortical parenchyma and the phloem. During retting, a number of morphological changes occur, mainly in the outer tissues of the stem (Figure 2A). Parenchyma cells around and between fiber bundles that can be easily observed during the early stages of retting almost disappear at later stages (Chabbert et al., 2020). Fiber bundles themselves also become progressively destructured mainly due to the dissociation of individual fiber cells (Akin, 2003; Figure 2A). Individual fiber cells are integrated into stem tissues via their compound middle lamella that not only links them to other fibers, but also links the peripheral bundle cells to the cortical cells in the external stem tissues (Meijer et al., 1995; Akin, 2013; Figure 2B). Retting therefore involves the progressive enzymatic degradation of both cortical parenchyma cell walls and the fiber compound middle lamella. Chemically these structures are composed of different polymers. Cellulose, hemicellulose (mainly xyloglucan), pectic polysaccharides (homogalacturonan, HG), rhamnogalacturonan I (RG-I), and rhamnogalacturonan II (RG-II), as well as some structural proteins are found in the primary cell wall. In contrast, the middle lamella is mainly composed of pectins forming what is commonly called the cellular cement. Fibers cells, in addition to a middle lamella and primary cell wall, have thick secondary walls that are rich in cellulose (up to 80%) and contain non-cellulosic polysaccharides such as galactans and glucomannans (Morvan et al., 2003; Rihouey et al., 2017). Low levels of lignin (2–5%) can also be sometimes found in the middle lamella, primary cell wall, and S1 layer of the secondary cell wall (Day et al., 2005). The technical properties of fibers are due to a high content of crystalline cellulose, with microfibrils almost parallel to the main axis of the cells (Müller et al., 1998). The Figure 2B shows the general organization of the plant cell wall.
Retting Enzymes
The degradation of cell wall polymers necessitates the intervention of enzymes belonging to a wide range of different enzyme families (Figure 2C). In recent studies on hemp and flax retting, exopolygalacturonase and α-L-arabinosidase, β-D-xylosidase and β-D-galactosidase, β-D-glucosidase and cellobiohydrolase activities were followed during the retting process (Bleuze et al., 2018; Chabbert et al., 2020). Pectin-degrading enzymes are distributed across nine families of carbohydrate-active enzymes (CAZymes), CE8, PL1, PL2, PL3, PL9, PL10, GH28, GH78, and GH88. GH28 polygalacturonases (PGAs) play an important role in the breakdown of pectins in pathogenic ‘fungi.’ Pectin esterases (family CE8) catalyze the de-esterification of pectins into pectate and methanol (Zhao et al., 2013, 2014). Several strains of filamentous ‘fungi,’ isolated from flax stems retted in the field, such as Rhizomucor pusillus (Mucoromycota formerly Zygomycota) and Fusarium lateritium (Ascomycota) have shown a capacity for decohesion of fibers, in particular thanks to a high level of pectinolytic activity (Henriksson et al., 1997). Pectinolytic activities (e.g., polygalacturonases or pectin lyases) have also been observed in various microorganisms during water-retting (Zhao et al., 2016). For example, Clostridium felsineum shows a strong pectinolytic activity and a good capacity for retting. Studies using enzymatic retting have shown that the use of polygalacturonases alone is sufficient for the decohesion of fibers (Evans et al., 2002; Akin et al., 2004). Some microorganisms used during retting such as Bacillus subtilis and Erwinia carotovora (syn. Pectobacterium carotovorum) are known to produce pectin lyases (Sharma, 1986). Saprophytic or parasitic microorganisms have CAZymes showing activities of hemicellulolytic types (vanden Wymelenberg et al., 2010). These activities, in particular those of xylanases, have been observed during water-retting (Donaghy et al., 1990), or in bacterial cultures resulting from the cultivation of dew retted flax stems (Sharma, 1986). Very recently, a study on retting with water in a fermenter, showed a cyclic evolution of mannanases, another hemicellulolytic enzyme (Zhao et al., 2016).
During retting the enzymatic degradation of cell wall polymers can be followed by monitoring the sugars released. For example, the release of fibers was correlated with a decrease in the concentration of galacturonic acid resulting from the breakdown of pectic material in the middle lamella at the end of retting (Rosemberg and de França, 1967). A recent study of retting dynamics by both scanning electron microscope (SEM) imaging and the analysis of certain wall polymers also confirmed the link between the degradation of primary cell walls and fiber middle lamella and the dissociation of fiber bundles (Chabbert et al., 2020). In contrast glucose and mannose/galactose levels remain stable suggesting that secondary cell wall polysaccharides are not degraded (Chabbert et al., 2020). The release of other cell wall polymers such as cutin, wax, and aromatic compounds (sinapyl alcohol and ferulic acid) has also been investigated by chemical analyzes and mass spectrometry in a comparative study of water- and field-retting (Morrison et al., 2000).
Factors Affecting Retting
At the plant level, the amount of pectins varies depending on the cultivar and will influence the duration of retting (Brown et al., 1986; Haag et al., 2017). Increasing maturity of the stems will also favor lignin deposition in the fiber middle lamella associated with difficulties in retting (Meijer et al., 1995; Pallesen, 1996). However, the influence of exogenous factors, either abiotic or biotic, also has an important impact on retting. In the first category, can be classified all the pedoclimatic factors such as the soil structure or the mineral elements (for example nitrogen, phosphorus and potassium), the physico-chemistry of the soil, the rotation of the cultures, the thickness of the swathe, as well as climatic conditions and seasonal variations (Sharma and Faughey, 1999). In the second category, the most influential factor will undoubtedly be the microbiota present in the soil and in/on the stem at the beginning of retting (Djemiel et al., 2017; Chabbert et al., 2020). Intriguingly, a recent study has revealed the existence of a very close link between the microbiome and the spatial and temporal development of the plant (composition, physiology), or even cultural conditions (Comeau et al., 2020). In the light of such observations it seems likely that plant growth and fiber development in flax will also be affected by the microbiome present. Furthermore, the microbiome itself will be influenced by plant growth and retting conditions. A more integrated approach is clearly necessary if we are to fully understand all of the factors affecting fiber quality (Figure 1).
Microbiology of Retting, What Have We Learned From Classical Approaches?
Since it is the microorganisms that produce the hydrolytic enzymes responsible for retting, an important step in our understanding of this process is to identify the organisms responsible. Historically, various bacteria and fungi were identified in a number of different studies using isolation and culture-based approaches. However, such strategies are not powerful enough to obtain a complete inventory of the microorganisms present as only a small percentage of taxa can be successfully cultured under laboratory conditions. Moreover, these approaches are generally inappropriate for dynamic studies looking at how microbial communities evolve during retting. More recently, microbial retting studies have greatly benefited from the use of HTS technologies that have produced exhaustive inventories of bacteria and fungi linked with this process. A list of microorganisms identified by classical methods is given in Tables 1, 2 and those identified by metabarcoding approaches in Table 3.
| Retting Mode | Phylum | Current Name (Name in Publication) | Localization | Method | References | |
| Water | Dew | |||||
| X | Firmicutes | Bacillus amylobacter | Not mentioned probably France | Not mentioned | van Tieghem (1879) | |
| X | Proteobacteria | Enterobacter aerogenes (Bacterium aerogenes) | Not mentioned | Screening on agar media or liquid culture | Allen (1946b) | |
| X | Proteobacteria | Escherichia coli (Bacterium coli) | Not mentioned | |||
| X | Proteobacteria | Streptococcus (Streptococci) | Not mentioned | |||
| X | Firmicutes | Lactobacillus (Lactobacilli) | Not mentioned | |||
| X | Firmicutes | Clostridium tertium | Not mentioned | Screening on agar media or liquid culture | Allen (1946a) | |
| X | Firmicutes | Clostridium | Australia (Melbourne) | Not mentioned | Lanigan (1950) | |
| X | Firmicutes | Clostridium felsineum | Australia (Melbourne) | |||
| X | Proteobacteria | Achromobacter parvulus | Brazil (Santa Catarina) | Screening on agar media culture | Rosemberg (1965) | |
| X | Proteobacteria | Aerobacter cloacae | Brazil (Santa Catarina) | |||
| X | Proteobacteria | Aerobacter aerogenes | Brazil (Santa Catarina) | |||
| X | Firmicutes | Bacillus brevis | Brazil (Santa Catarina) | |||
| X | Firmicutes | Bacillus cereus | Brazil (Santa Catarina) | |||
| X | Firmicutes | Bacillus megaterium | Brazil (Santa Catarina) | |||
| X | Firmicutes | Bacillus sphaericus | Brazil (Santa Catarina) | |||
| X | Firmicutes | Bacillus subtilis | Brazil (Santa Catarina) | |||
| X | Firmicutes | Clostridium butylicum | Brazil (Santa Catarina) | |||
| X | Firmicutes | Clostridium beijerinckii | Brazil (Santa Catarina) | |||
| X | Firmicutes | Clostridium saprogenes | Brazil (Santa Catarina) | |||
| X | Firmicutes | Clostridium sartagoformum | Brazil (Santa Catarina) | |||
| X | Firmicutes | Clostridium saccharoacetoperbutylicum | Brazil (Santa Catarina) | |||
| X | Firmicutes | Clostridium perenne | Brazil (Santa Catarina) | |||
| X | Proteobacteria | Escherichia coli | Brazil (Santa Catarina) | |||
| X | Actinobacteria | Gaffkya tetragena (probably contamination) | Brazil (Santa Catarina) | |||
| X | Proteobacteria | Pseudomonas aeruginosa | Brazil (Santa Catarina) | |||
| X | Proteobacteria | Pseudomonas pseudomallei | Brazil (Santa Catarina) | |||
| X | Proteobacteria | Paracolobactrum aerogenoides | Brazil (Santa Catarina) | |||
| X | Proteobacteria | Serratia plymuthica | Brazil (Santa Catarina) | |||
| X | Firmicutes | Staphylococcus epidermis (probably contamination) | Brazil (Santa Catarina) | |||
| X | Firmicutes | Bacillus mycoides | Northern Ireland (Lambeg) | Screening on agar media culture | Sharma (1986) | |
| X | Firmicutes | Bacillus subtilis | Northern Ireland (Lambeg) | |||
| X | Proteobacteria | Erwinia carotovora | Northern Ireland (Lambeg) | |||
| X | Proteobacteria | Pseudomonas fluorescens | Northern Ireland (Lambeg) | |||
| X | Proteobacteria | Pseudomonas putida | Northern Ireland (Lambeg) | |||
| X | Actinobacteria | Micrococcus sp. | Northern Ireland (Lambeg) | |||
| X | Firmicutes | Bacillus subtilis | Northern Ireland (Lambeg) | Screening on agar media culture | Donaghy et al. (1990) | |
| X | Actinobacteria | Cellulomonas spp. | Northern Ireland (Lambeg) | |||
| X | Firmicutes | Clostridium felsineum | Northern Ireland (Lambeg) | |||
| X | Firmicutes | Bacillus cereus | Northern Ireland (Lambeg) | |||
| X | Firmicutes | Clostridium felsineum | Italy | Screening on agar media or liquid culture and identification with 16S rDNA | Tamburini et al. (2003) | |
| X | Firmicutes | Anaerobacter polyendosporus | Italy | |||
| X | Firmicutes | Clostridium saccharobutylicum | Italy | |||
| X | Firmicutes | Clostridium aurantibutyricum | Italy | |||
| X | Firmicutes | Clostridium acetobutylicum | Italy | |||
| X | Firmicutes | Bacillus subtilis | Italy | |||
| X | Firmicutes | Bacillus pumilus | Italy | |||
| X | Firmicutes | Paenibacillus amylolyticus | Italy | |||
| Retting Mode | Phylum | Current Name (Name in Publication) | Localization | Method | References | ||
| Water | Dew | Standing* | |||||
| X | Ascomycota | Epicoccum nigrum | Ireland (Lambeg, Hillsborough) | Screening on agar media culture | Brown (1984) | ||
| X | Mucoromycota (Zygomycota) | Rhizopus sp. | |||||
| X | Mucoromycota (Zygomycota) | Mucor sp. | |||||
| X | Ascomycota | Cladosporium herbarum | |||||
| X | Ascomycota | Botrytis cinerea | |||||
| X | Ascomycota | Penicillium sp. | |||||
| X | Ascomycota | Fusarium culmorum | |||||
| X | Ascomycota | Phoma sp. | |||||
| X | Ascomycota | Alternaria spp. | |||||
| X | Ascomycota | Yeasts | |||||
| X | Ascomycota | Cladosporium herbarum | Northern Ireland (Lambeg) | Screening on agar media culture | Sharma (1986) | ||
| X | Ascomycota | Fusarium culmorum | |||||
| X | Ascomycota | Botrytis cinerea | |||||
| X | Ascomycota | Epicoccum nigrum | |||||
| X | Ascomycota | Yeast | |||||
| X | Ascomycota | Alternaria spp. | |||||
| X | Ascomycota | Aspergillus flavus | Italy (Budrio) | Not mentioned | Fila et al. (2001) | ||
| X | Ascomycota | Aspergillus niger | |||||
| X | Ascomycota | Epicoccum nigrum | |||||
| X | Ascomycota | Fusarium oxysporum | |||||
| X | Mucoromycota (Zygomycota) | Mucor hiemalis | |||||
| X | Ascomycota | Penicillium simplicissimum | |||||
| X | Mucoromycota (Zygomycota) | Rhizopus stolonifer | |||||
| X | Ascomycota | Fusarium equiseti | United States (South Carolina) | Screening on agar media culture | Henriksson et al. (1997) | ||
| X | Ascomycota | Yeast | |||||
| X | Mucoromycota (Zygomycota) | Rhizomucor pusillus | |||||
| X | Ascomycota | Trichoderma virens | |||||
| X | Ascomycota | Alternaria alternata | |||||
| X | Ascomycota | Fusarium lateritium | United States (Connecticut) | ||||
| X | Ascomycota | Cladosporium herbarum | |||||
| X | Ascomycota | Fusarium oxysporum | France | ||||
| X | Ascomycota | Epicoccum nigrum | Holland | ||||
| Plant Fiber | Retting Types | Microbes Studied | Molecular Marker Used | Sequencer | Years | OTU Richness (mean) | Subsampling | Major Phyla | References |
| Kenaf (Hibiscus cannabinus) | Water | Bacteria | 16S rRNA | Ion torrent PGM | 2013 | 1,500 | 28,000 | Firmicutes, Proteobacteria, Bacteroidetes | Visi et al. (2013) |
| Flax (Linum usitatissimum) | Water | Bacteria | 16S rRNA | Illumina MiSeq | 2016 | 70 | NM | Bacteroidetes, Firmicutes, Proteobacteria | Zhao et al. (2016) |
| Flax (Linum usitatissimum) | Dew | Bacteria; Fungi | 16S rRNA; ITS2 | Illumina MiSeq | 2017 | 300; 220 | 20,548; 42,436 | Proteobacteria, Bacteroidetes, Actinobacteria, Firmicutes; Ascomycota, Basidiomycota | Djemiel et al. (2017) |
| Flax (Linum usitatissimum) | Dew | Bacteria; Fungi | 16S rRNA; ITS2 | Illumina MiSeq | 2020 | 200; 260 | 5,919; 86,050 | Proteobacteria, Actinobacteria, Bacteroidetes; Ascomycota, Basidiomycota | Chabbert et al. (2020) |
| Hemp (Cannabis sativa) | Greenhouse | Bacteria | 16S rRNA | Illumina MiSeq | 2020 | 100 | 2,234 | Proteobacteria, Bacteroidetes | Law et al. (2020) |
| Kenaf (Hibiscus cannabinus) | Water | Bacteria; Fungi | 16S rRNA; 18S rRNA | Illumina MiSeq | 2020 | 430* | NM | Bacteroidetes, Proteobacteria; Basidiomycota, Ascomycota | Duan et al. (2020) |
Bacteria Identified
The first reported study focusing on the retting of flax with water was carried out by the French biologist and botanist Philippe Édouard Léon Van Tieghem (1839–1914), who concluded that the bacteria Bacillus amylobacter was probably responsible for the decomposition of pectins (Figure 3A; van Tieghem, 1879). The second genus to be associated with water-retting was the genus Clostridium and more particularly Clostridium felsineum (Figure 3A; Lanigan, 1950). A few years later, Rosemberg (1965) isolated 22 individuals including species of the Clostridium and Pseudomonas genera. Pseudomonas aeruginosa was recognized as the fastest species in fiber decohesion (Rosemberg, 1965). This study also identified, for the first time, Achromobacter parvulus which appears to be involved in the latter stages of retting.
Later, Sharma (1986) isolated and identified six bacterial species from the Actinobacteria, Firmicutes, and Proteobacteria phyla as well as five fungal species belonging to the Ascomycota phylum (Figure 3B).
In 1990, a study at the University of Ulster (Northern Ireland) followed the evolution of anaerobic bacteria during water-retting, both at an industrial and laboratory scale. Bacillus licheniformis and Bacillus subtilis were the most dominant during the first and second phase, corresponding to the growth phase and the rapid pectinolytic phase (between 10–40 h) of water-retting. Clostridium acetobutylicurn and Clostridium felsineum appear during the last phase known as the slow pectinolytic phase (Figure 3A; Donaghy et al., 1990). All the bacteria considered belong to the Firmicutes phylum.
In 2003, another study provided further information about pectinolytic bacteria involved in water-retting. Although this study still involved a culture step to isolate bacteria, it was the first one to use a molecular marker approach, in this case by amplifying a partial region of the 16S rRNA gene, using the (amplified ribosomal DNA restriction analysis) ARDRA technique. All of the anaerobic strains were assigned to the Clostridium genus and the aerobic strains to the Bacillus or Paenibacillus genera. Anaerobic colonies with significant polygalacturonase activity belonged to two phylogenetic clusters assigned to the Clostridium acetobutylicum/Clostridium felsineum and Clostridium saccharobutylicum species. For aerobic bacteria, colonies with significant polygalacturonase activity belonged to two phylogenetic clusters assigned to the Bacillus subtilis species (Figure 3A; Tamburini et al., 2003). All these strains also belong to the Firmicutes phylum.
Fungi Identified
One of the first studies on the microbiology of dew-retting identified several genera and species from the Fungi kingdom belonging to the Ascomycota and Mucoromycota (formerly Zygomycota) phyla (Figure 3B; Brown, 1984). In the late 1990s, an American study looked at fungi involved in flax dew-retting in the United States, Netherlands, and France (Figure 3B; Henriksson et al., 1999). Seven strains of filamentous fungi (including six still not described in the literature to this day) and a yeast, could be identified following isolation, cultivation on synthetic media, and purification in order to compare their activities and their efficiency in fiber release. Another study involving in vitro retting tests was carried out by Fila and coworkers but no new fungal genus was identified (Figure 3B; Fila et al., 2001).
Some fungi have been characterized as over-retting actors, for example Fusarium lateritium and more particularly Epicoccum nigrum, recognized as being a primary saprophyte in retting (Brown, 1984; Henriksson et al., 1997; Akin et al., 1998). Another species, Rhizomucor pusillus has been observed to degrade part of the surface of the cuticle, thus probably allowing the entry of microorganisms (Henriksson et al., 1997).
The overall conclusion that can be drawn from these classical, culture-based studies of retting is that the type of microorganism identified depends heavily upon the retting type. Field retting is an aerobic environment and although both bacteria and fungi are identified it is generally the latter that predominate with species belonging mostly to the Ascomycota phylum and to a lesser extent the Mucoromycota phylum (formerly Zygomycota; Table 2). In contrast, water-retting mainly involves anaerobic microorganisms (bacteria). Nevertheless, several fungi have also been identified in various anaerobic environments and classified within two very close phyla, Neocallimastigomycota and Chytridiomycota (Griffith et al., 2010; Gruninger et al., 2014). Interestingly, some of these anaerobic fungi, are also found in the rumen, and have been shown to possess a number of GH genes probably resulting from horizontal transfers of bacterial genes (Garcia-Vallvé et al., 2000; Steenbakkers et al., 2001).
Metabarcoding and the Breakthrough in Knowledge on Retting Microbial Diversity
Metabarcoding, also known as targeted metagenomics, is an approach that has revolutionized the study of microbial communities. In recent years the decrease in sequencing costs has allowed it to become more democratic, leading to the use of this approach in a larger number of laboratories and studies. The large and growing number of publications that use metabarcoding reflects this evolution regardless of the habitat explored, including studies on the microbiota of plants (Figure 4). Metabarcoding consists in sequencing an amplicon (most often corresponding to one or more regions of a phylogenetic marker) on a high−throughput platform to characterize microbial diversity from complex/environmental samples, thereby enabling studies of alpha-diversity, community structure or taxonomy (Deiner et al., 2017; Piper et al., 2019; Tedersoo et al., 2019).
There are currently very few studies on the holistic interactions between microorganisms and the flax plant during its growth and their implication on the quality of fibers. The only interactions studied for the moment are those taking place during retting, and that concern the capacity of microorganisms to degrade the plant cell wall and therefore in fine to affect fiber quality for their future use (Table 3). The first investigation using the metabarcoding approach investigated bacterial communities during kenaf water-retting under different controlled conditions (Visi et al., 2013). The authors used the 16S rRNA gene as the molecular marker and performed sequencing with an Ion Torrent Personal Genome Machine (PGM) system. The operational taxonomic unit (OTU) richness values from Visi et al. (2013) seem to be extremely high compared to other water-retting studies (Table 3). More recently, the evolution of both bacterial and fungal communities during water-retting of kenaf was studied using 18S rRNA gene (Duan et al., 2020). However, the results obtained concerning the Fungi kingdom must be taken with care because precisions concerning the bioinformatics analysis are not available (e.g., sub-sampling size/rarefaction) and certain inaccuracies on the taxonomic affiliation occured (e.g., confusion between the different taxonomic levels; Opisthokonta has been classified as a subkingdom and a phylum). To date, only one study has been conducted on hemp retting by metabarcoding (Law et al., 2020). This study indicated an average OTU richness mean of one hundred. In comparison, the study by Ribeiro et al. (2015) identified 66 different bacterial phylotypes. Despite the fact that these two studies involved different environmental conditions –Law et al. (2020) conducted hemp retting in a greenhouse under controlled conditions, while Ribeiro et al. (2015) collected samples in the field – the OTU or phylotype distributions were similar for the Proteobacteria (85 vs. 77%), Bacteroidetes (8 vs. 11%) and Firmicute (1% or less) but differed for Actinobacteria (2.3 vs. 11%).
Until now, it is flax retting that has been the most studied by metabarcoding, with two studies conducted on field retting (Djemiel et al., 2017) and one on water-retting in tanks (Zhao et al., 2016; Figure 3A). Zhao et al. (2016) focused on the evolution of anaerobic bacteria using a partial 16S rRNA gene sequence region following the addition of a strain of Bacillus cereus HDYM-02, previously characterized in their laboratory as being very effective in the decohesion of fibers. Interestingly, this species had been identified previously as an efficient retting agent that did adversely affect fiber mechanical properties (Donaghy et al., 1990). The study by Zhao et al. (2016) demonstrated the power of HTS since they were able to identify five phyla (Actinobacteria, Cyanobacteria, Bacteroidetes, Firmicutes, and Proteobacteria) and eight classes (Flavobacteriia, Sphingobacteriia, Bacilli, Clostridia, Negativicutes, α-proteobacteria, β-proteobacteria, and γ-proteobacteria), some of which were not previously known to be involved in water-retting (e.g., Cyanobacteria and Bacteroidetes for phyla, and Flavobacteriia, Sphingobacteriia, Negativicutes, α-proteobacteria, and β-proteobacteria for classes; Zhao et al., 2016).
The interest of the metabarcoding approach for flax field-retting was also demonstrated by two recent studies (Djemiel et al., 2017; Chabbert et al., 2020). The first study explored the diversity and dynamics of bacterial and fungal communities involved in this process by using an HTS DNA metabarcoding approach (16S rRNA/Internal Transcribed Spacer (ITS) region, Illumina Miseq) on plant and soil samples collected over a period of 7 weeks in July and August 2014 (Djemiel et al., 2017). Twenty-three bacterial and six fungal phyla were identified in soil samples and 11 bacterial and four fungal phyla in plant samples. Dominant phyla identified were Proteobacteria, Bacteroidetes, Actinobacteria, and Firmicutes (bacteria) and Ascomycota, Basidiomycota, and Zygomycota (fungi) all of which have been previously associated with flax dew-retting except for Bacteroidetes and Basidiomycota that were identified for the first time. The use of this powerful method also allowed the identification of rare phyla never previously associated with retting: Acidobacteria, CKC4, Chlorobi, Fibrobacteres, Gemmatimonadetes, Nitrospirae, and TM6 (bacteria), and Chytridiomycota (fungi). These results perfectly illustrate the efficiency of metabarcoding for detecting phyla of low abundance. In addition, this study also revealed that the agricultural practice of swathe turning affected bacterial and fungal community structure, and most likely contributed to a more uniform retting. The use of the prediction tool Phylogenetic Investigation of Communities by Reconstruction of Unobserved States (PICRUSt) (Langille et al., 2013; Douglas et al., 2020); was also used on OTU tables to decipher the functional potential of bacterial communities. This analysis predicted a large collection of potential bacterial enzymes capable of hydrolyzing the backbones and side-chains of cell wall polysaccharides. These results demonstrate the interest of a combined metabarcoding and PICRUSt analysis to predict functional composition of bacterial communities. A similar approach reported by Chabbert et al. (2020) allowed a comparison of the evolution of bacterial and fungal retting communities under different environmental conditions. Eight out of the ten most abundant bacterial OTUs identified in this study were also among the top ten OTUs identified in the 2014 study by Djemiel et al. (2017) in the same geographical area, but in a different field and under different climatic conditions. In contrast, only half of the top ten fungal OTUs were retrieved. In this last study, enzyme activities were measured in addition to targeted metagenomics showing the presence of a high microbial diversity with potential enzymatic functions during retting. The comparison of these two studies could suggest that a year-to-year variability exists, perhaps more so in fungal communities compared to bacteria. Nevertheless, the identification of comparable enzymatic functions in the two studies would suggest a functional redundancy within the communities. In conclusion, the contribution of metabarcoding to the knowledge about the microbiology of retting is undeniable. In addition, the technique can also be adapted to ask questions about the living world in the widest possible way. For example, by refining the choice of primers metabarcoding can also be used to inventory often neglected organisms such as Archaea or microeukaryotes (Bahram et al., 2019).
Concluding Remarks and Future Perspectives
A multitude of intrinsic and extrinsic factors, both during plant growth and after harvest, all contribute to establishing the final phenotype of plant fibers reaching the factory (Figure 1). Since these factors affect fiber morphology, cell wall composition and organization they will also impact on the behavior of the separated fibers during subsequent industrial processing. In this paper we have focused on the different meta-omic technologies that have been, or could be, exploited to improve our understanding of one of these many factors - the field retting step of flax. This step involves a complex interaction between the harvested plant material and microorganisms, both in the soil and on the plant at the time of harvest. However, the complexity of this interaction does not stop there since the nature of the plant material itself (e.g., fiber morphology and cell wall composition), as well as the composition and functional capacity of the microbiome are also impacted by genetics and environmental conditions. In the light of such observations it is becoming clear that not only do we need to implement a multi-omics systems biology approach, but that we also need to take into account all of the actors that contribute to producing the final phenotype. This idea is embodied in the concept of the ‘holobiont’ which states that we should no longer consider the plant as an isolated ‘stand-alone’ organism, but rather as an individual with its associated microbial communities and in which their overall interactions are modulated by the pedo-climatic environment (Simon et al., 2019). Their association plays a decisive role in terms of the biodiversity and functionality of this ecosystem in which all the partners influence each other. Moreover, plant/microorganism (symbiotic, pathogenic, saprophytic) and microorganism/microorganism interactions will be affected by different biotic factors (e.g., species, cultivar, age, health, and stages of plant development) and abiotic factors (e.g., soil physicochemical composition, climatic conditions (Comeau et al., 2020; Fitzpatrick et al., 2020). As demonstrated in this review the combination of related ‘-omics’ such as metatranscriptomics and metaproteomics, together with biochemical studies, offers the unique opportunity to investigate the particular holobiont constituted by the overall fiber plant growing cycle. There is no doubt that the comprehension of this complex ecological machinery promises to make a major contribution to the control of plant natural fiber quality. For this purpose, it will be necessary to integrate together large multiscale data sets consisting of fiber parameters, biological and environmental information with the development of predictive models to be processed by AI analyses. The collection of massive data would rely on the development of connected microsensors reporting different parameters in real time directly from the field. Flax farmers will then have at their disposal a ‘retting toolbox’ making use of molecular markers, metabolite, protein and gene expression, biochemistry and phenotyping (morphological and agronomic) data for targeting selected traits. The cultivation of flax will then enter the era of smart connected agriculture.
Conflict of Interest
The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.