Gene Disruption Method Research Articles

Efficient multiple gene knockout in Colletotrichum higginsianum via CRISPR/Cas9 ribonucleoprotein and URA3-based marker recycling.

Colletotrichum higginsianum is a hemibiotrophic pathogen that causes anthracnose disease on crucifer hosts, including Arabidopsis thaliana. Despite the availability of genomic and transcriptomic information and the ability to transform both organisms, identifying C. higginsianum genes involved in virulence has been challenging due to recalcitrance to gene targeting and redundancy of virulence factors. To overcome these obstacles, we developed an efficient method for multiple gene disruption in C. higginsianum by combining CRISPR/Cas9 and a URA3-based marker recycling system. Our method significantly increased the efficiency of gene knockout via homologous recombination by introducing genomic DNA double-strand breaks. We demonstrated the applicability of the URA3-based marker recycling system for multiple gene targeting in the same strain. Using our technology, we successfully targeted two melanin biosynthesis genes, SCD1 and PKS1, which resulted in deficiency in melanization and loss of pathogenicity in the mutants. Our findings demonstrate the effectiveness of our methods in analysing virulence factors in C. higginsianum, thus accelerating research on plant-fungus interactions.

Introduction of a CRISPR-nCas9 Gene-Targeting Plasmid into Staphylococcus aureus for Gene Disruption.

Methods for gene disruption are essential for functional genomics, and there are multiple approaches for altering gene function in bacteria. One of these methods involves introducing a premature stop codon in a gene of interest, which can be achieved by using the CRISPR-nCas9-cytidine deaminase system. The approach involves the mutation of editable cytidines to thymidines, with the goal of generating a novel stop codon that ultimately results in a nonfunctional gene product. The workflow involves two major sections, one for the identification of editable cytidines, the design of the targeting spacer oligonucleotides for introduction into the CRISPR-nCas9 cytidine deaminase plasmid, and the construction of the gene-targeting CRISPR-nCas9 cytosine deaminase plasmids, and one for the actual introduction of the mutation in the species of interest. Here, we describe the steps for the second part. Specifically, we describe (1) how to introduce the gene-targeting pnCasSA-BEC plasmid into Staphylococcus aureus, (2) how the gene inactivation in S. aureus can be confirmed by PCR and sequencing, and (3) how, following successful gene inactivation, the strain can be cured of the pnCasSA-BEC plasmid. To better illustrate the method, and as specific example, two different geh gene-inactivation mutations are generated here in S. aureus RN4220. The protocol, however, can easily be adapted to generate other gene-inactivating mutations.

Identification of Editable Sites, Spacer Oligonucleotide Design, and Generation of the Gene-Targeting CRISPR-nCas9 Plasmid for Gene Disruption in Staphylococcus aureus Using the CRISPR-nCas9 and Cytidine Deaminase System.

Methods for gene disruption are essential for functional genomics, and there are multiple approaches for altering gene function in bacteria. One of these methods involves introducing a premature stop codon in a gene of interest, which can be achieved by using the CRISPR-nCas9-cytidine deaminase system. The approach involves the mutation of editable cytidines to thymidines, with the goal of generating a novel stop codon that ultimately results in a nonfunctional gene product. The workflow involves two major sections, one for the identification of editable cytidines, the design of the targeting spacer oligonucleotides for introduction into the CRISPR-nCas9 cytidine deaminase plasmid, and the construction of the gene-targeting CRISPR-nCas9 cytosine deaminase plasmids, and one for the actual introduction of the mutation in the species of interest. Here, we describe the steps for the first part. To better illustrate the method and oligonucleotide design, we describe the construction of Staphylococcus aureus RN4220 geh mutants with C to T base changes at two different positions, leading to the construction of strains RN4220-geh(160stop) and RN4220-geh(712stop). We outline the steps for (1) the identification of editable cytidines within genes using the CRISPR-CBEI toolkit website, and (2) the design of the targeting spacer oligonucleotides for introduction into the CRISPR-nCas9 cytidine deaminase plasmid pnCasSA-BEC, followed by (3) the construction of the gene-targeting (in this example, geh gene-targeting) CRISPR-nCas9 cytosine deaminase plasmids pnCasSA-BEC-gehC160T and pnCasSA-BEC-gehC712T using the Golden Gate assembly method, plasmid recovery in Escherichia coli, and confirmation by colony PCR and sequencing. The method can be easily adapted to construct gene-inactivation mutants in other S. aureus genes.

Strain breeding and gene complementation strategies for filamentous fungi

Filamentous fungi produce a variety of unique bioactive metabolites, which are important to food, medical and agricultural industries. In order to better utilize filamentous fungi for synthesizing their valuable products, platform technologies, which include gene disruption and complementation methods, are increasingly expected for editing and analyzing gene functions involved in their metabolic pathways. As more and more genomes of filamentous fungi have been sequenced and the efficient clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 system has been widely utilized for the genetic manipulation of filamentous fungi, gene modification and gene function analyses become more feasible. However, gene function studies on some species of filamentous fungi are still limited by gene editing or complemented strategies. In this review, we summarized the current strain breeding methods, gene complementation strategies and biotechnological application of gene complementation for filamentous fungi. In addition, we discuss the challenges to current gene complementation for biotechnological application and show perspectives on its future technological development and application to biotechnology studies on filamentous fungi.

Development of Efficient Genome-Reduction Tool Based on Cre/loxP System in Rhodococcus erythropolis.

Rhodococcus has been extensively studied for its excellent ability to degrade artificial chemicals and its capability to synthesize biosurfactants and antibiotics. In recent years, studies have attempted to use Rhodococcus as a gene expression host. Various genetic tools, such as plasmid vectors, transposon mutagenesis, and gene disruption methods have been developed for use in Rhodococcus; however, no effective method has been reported for performing large-size genome reduction. Therefore, the present study developed an effective plasmid-curing method using the levansucrase-encoding sacB gene and a simple two-step genome-reduction method using a modified Cre/loxP system. For the results, R. erythropolis JCM 2895 was used as the model; a mutant strain that cured all four plasmids and deleted seven chromosomal regions was successfully obtained in this study. The total DNA deletion size was >600 kb, which corresponds mostly to 10% of the genome size. Using this method, a genome-structure-stabilized and unfavorable gene/function-lacking host strain can be created in Rhodococcus. This genetic tool will help develop and improve Rhodococcus strains for various industrial and environmental applications.

Development of the CRISPR-Cas9 System for the Marine-Derived Fungi Spiromastix sp. SCSIO F190 and Aspergillus sp. SCSIO SX7S7.

Marine-derived fungi are emerging as attractive producers of structurally novel secondary metabolites with diverse bioactivities. However, the lack of efficient genetic tools limits the discovery of novel compounds and the elucidation of biosynthesis mechanisms. Here, we firstly established an effective PEG-mediated chemical transformation system for protoplasts in two marine-derived fungi, Spiromastix sp. SCSIO F190 and Aspergillus sp. SCSIO SX7S7. Next, we developed a simple and versatile CRISPR-Cas9-based gene disruption strategy by transforming a target fungus with a single plasmid. We found that the transformation with a circular plasmid encoding cas9, a single-guide RNA (sgRNA), and a selectable marker resulted in a high frequency of targeted and insertional gene mutations in both marine-derived fungal strains. In addition, the histone deacetylase gene rpd3 was mutated using the established CRISPR-Cas9 system, thereby activating novel secondary metabolites that were not produced in the wild-type strain. Taken together, a versatile CRISPR-Cas9-based gene disruption method was established, which will promote the discovery of novel natural products and further biological studies.

A streamlined strain engineering workflow with genome-wide screening detects enhanced protein secretion in Komagataella phaffii.

Expression of secreted recombinant proteins burdens the protein secretion machinery, limiting production. Here, we describe an approach to improving protein production by the non-conventional yeast Komagataella phaffii comprised of genome-wide screening for effective gene disruptions, combining them in a single strain, and recovering growth reduction by adaptive evolution. For the screen, we designed a multiwell-formatted, streamlined workflow to high-throughput assay of secretion of a single-chain small antibody, which is cumbersome to detect but serves as a good model of proteins that are difficult to secrete. Using the consolidated screening system, we evaluated >19,000 mutant strains from a mutant library prepared by a modified random gene-disruption method, and identified six factors for which disruption led to increased antibody production. We then combined the disruptions, up to quadruple gene knockouts, which appeared to contribute independently, in a single strain and observed an additive effect. Target protein and promoter were basically interchangeable for the effects of knockout genes screened. We finally used adaptive evolution to recover reduced cell growth by multiple gene knockouts and examine the possibility for further enhancing protein secretion. Our successful, three-part approach holds promise as a method for improving protein production by non-conventional microorganisms.

A modified method of gene disruption in Komagataella phaffii with Cre/loxP system

To increase protein production, technologies of gene manipulation for engineering the yeast Komagataella phaffii are extensively exploited. In this study, we developed a convenient gene disruption method in the yeast via Cre/loxP system. First, the simple gene disruption cassette [upstream homologous region (UP)-lox71-Sh ble-lox66-downstream homologous region (DW)] was constructed and transformed into the yeast to replace target gene. Second, the Sh ble gene of the cassette integrated in the chromosome was inserted with the auxiliary plasmid pPICZαA/cre/his4, resulting in an expanded cassette of UP-lox71-Sh ble-pPICZαA/cre/his4-lox66-DW. The auxiliary plasmid was generated via sequential insertion of cre and his4 genes into pPICZαA, and linearized with SmaI before its transformation. Finally, for deletion of the sequence between lox71 and lox66 sites in the expanded cassette, CRE protein responsible for Cre/loxP-mediated recombination was produced by methanol induction. Consequently, the corresponding sequence was eliminated permanently, only leaving a scar of lox72 site in the disrupted genes. This strategy was verified by disrupting two genes in the yeast. As the markers were recycled, it was also suitable for multiple gene disruption.

Plasmid-Based Gene Knockout Strategy with Subsequent Marker Recycling in Pichia pastoris.

Gene knockout is a key technology in the development of cell factories and basic research alike. The methylotrophic yeast Pichia pastoris is typically employed as a producer of proteins and of fine chemicals, due to its ability to accumulate high cell densities in conjunction with a set of strong inducible promoters. However, protocols for genome engineering in this host are still cumbersome and time-consuming. Moreover, extensive genome engineering raises the need for a multitude of selection markers, which are limited in P. pastoris. In this chapter, we describe a fast and efficient method for gene disruption in P. pastoris that utilizes marker recycling to enable repetitive genome engineering cycles. A set of ready-to-use knockout vectors simplifies cloning procedures and facilitates quick knockout generation.

Candida albicans Secreted Aspartic Protease 7 is Essential for Damage of Human Oral Epithelial Cells

Aims: Candida albicans is an important human fungal pathogen in clinical settings. It possesses a wide spectrum of virulence traits, including but not limited to the production of Secreted Aspartic Proteases (SAPs), to invade host cells under predisposing conditions. The aims of the present study were to investigate the functional role of C. albicans SAP7 in invasion ability. Methods: The present study was carried out to construct C. albicans sap7Δ/Δ mutant strain using a PCR-based gene disruption method. The behaviors of this SAP7 knockout strain was evaluated and compared with the wild type and SAP7 complemented strains between human oral epithelial cells with respect to endocytosis, invasion, and tissue damage. Results: Compared with the wild type C. albicans strain, a 52% reduction in the endocytosis of the sap7Δ/Δ mutant strain by oral epithelial cells was observed, as well as a 25% attenuation of internalization, and a 27% reduction of tissue damage (P<0.05). Conclusion: Our data clearly demonstrates that C. albicans SAP7 contributes to tissue invasion into human oral epithelial cells which warrant further investigations as potential targets for antifungal interventions.

An efficient marker recycling system for sequential gene deletion in a deep sea-derived fungus Acremonium sp. HDN16-126

Acremonium species are prolific producers of therapeutic molecules which include the widely used beta-lactam antibiotic, cephalosporin. In light of their significant medical value, an efficient gene disruption method is required for the physiological and biochemical studies on this genus of fungi. However, the number of selection markers that can be used for gene targeting is limited, which constrain the genetic analysis of multiple functional genes. In this study, we established a uridine auxotrophy based marker recycling system which achieves scarless gene deletion, and allows the use of the same selection marker in successive transformations in a deep sea-derived fungus Acremonium sp. HDN16-126. We identified one homologue of Acremonium chrysogenum pyrG (also as a homologous gene of the yeast URA3) from HDN16-126, designated as pyrG-A1, which can be used as a selection marker on uridine free medium. We then removed pyrG-A1 from HDN16-126 genome via homologous recombination (HR) on MM medium with 5-fluoroortic acid (5-FOA), a chemical that can be converted into a toxin of 5-flurouracil by pyrG-A1 activity, thus generating the HDN16-126-△pyrG mutant strain which showed auxotrophy for uridine but insensitivity to 5-FOA and enabled the use of exogenous pyrG gene as both positive and negative selection marker to achieve the scarless deletion of target DNA fragments. We further applied this marker recycling system to successfully disrupt two target genes pepL (encodes a putative 2OG-Fe (II) dioxygenase) and pepM (encodes a putative aldolase) identified from HDN16-126 genome, which are proposed to be functional genes related to 2-aminoisobutyric acid metabolism in fungi. This work is the first application of uridine auxotrophy based scarless gene deletion method in Acremonium species and shows promising potential in assisting sequential genetic analysis of filamentous fungi.

CRISPR-Cas9 cytidine and adenosine base editing of splice-sites mediates highly-efficient disruption of proteins in primary and immortalized cells

CRISPR-Cas9 cytidine and adenosine base editors (CBEs and ABEs) can disrupt genes without introducing double-stranded breaks by inactivating splice sites (BE-splice) or by introducing premature stop (pmSTOP) codons. However, no in-depth comparison of these methods or a modular tool for designing BE-splice sgRNAs exists. To address these needs, we develop SpliceR (http://z.umn.edu/spliceR) to design and rank BE-splice sgRNAs for any Ensembl annotated genome, and compared disruption approaches in T cells using a screen against the TCR-CD3 MHC Class I immune synapse. Among the targeted genes, we find that targeting splice-donors is the most reliable disruption method, followed by targeting splice-acceptors, and introducing pmSTOPs. Further, the CBE BE4 is more effective for disruption than the ABE ABE7.10, however this disparity is eliminated by employing ABE8e. Collectively, we demonstrate a robust method for gene disruption, accompanied by a modular design tool that is of use to basic and translational researchers alike.

Candida albicans Bgl2p, Ecm33p, and Als1p proteins are involved in adhesion to saliva-coated hydroxyapatite

ABSTRACT Introduction: Candida albicans is an opportunistic pathogen that causes oral candidiasis. A previous study showed that Bgl2p and Ecm33p may mediate the interaction between the yeast and saliva-coated hydroxyapatite (SHA; a model for the tooth surface). This study investigated the roles of these cell wall proteins in the adherence of C. albicans to SHA beads. Methods: C. albicans BGL2 and ECM33 null mutants were generated from wild-type strain SC5314 by using the SAT1-flipper gene disruption method. A novel method based on labelling the yeast with Nile red, was used to investigate the adherence. Results: Adhesion of bgl2Δ and ecm33Δ null mutants to SHA beads was 76.4% and 64.8% of the wild-type strain, respectively. Interestingly, the adhesion of the bgl2Δ, ecm33Δ double mutant (87.7%) was higher than that of both single mutants. qRT-PCR analysis indicated that the ALS1 gene was over-expressed in the bgl2Δ, ecm33Δ strain. The triple null mutant showed a significantly reduced adherence to the beads, (37.6%), compared to the wild-type strain. Conclusion: Bgl2p and Ecm33p contributed to the interaction between C. albicans and SHA beads. Deletion of these genes triggered overexpression of the ALS1 gene in the bgl2Δ/ecm33Δ mutant strain, and deletion of all three genes caused a significant decrease in adhesion.

MaPmt1, a protein O-mannosyltransferase, contributes to virulence through governing the appressorium turgor pressure in Metarhizium acridum

O-glycosylation is a very important post-translational modification of protein and involved in many cell processes in fungi. There exist three protein O-manosyltransferanse genes (MaPmt1, MaPmt2, MaPmt4) in Metarhizium acridum based on sequence homology. Here, MaPmt1, a gene for Pmt1 O-manosyltransferanse in M. acridum, was characterized and functionally analyzed through targeted gene disruption and complementation methods. Deletion of MaPmt1 had no effect on conidial germination, but slightly increased the conidial yield and significantly impaired fungal tolerances to UV-B radiation and wet-heat. Deletion of MaPmt1 made the fungus become more sensitive to cell wall disturbing agents and exhibit a thinner cell wall with changed components. Insect bioassays showed that disruption of MaPmt1 attenuated the fungal virulence significantly by topical inoculation but not by injection, indicating that MaPmt1 is required for penetration during the infection of M. acridum. Interestingly, deletion of MaPmt1 did not affect appressorium formation but significantly decreased appressorium turgor pressure. Moreover, the decreased virulence of MaPmt1 disruptant is mainly due to the reduced appressorium turgor pressure, which may be resulted from the declined glycerol concentration, combined with the weakened cell wall that could not hold the normal appressorium turgor pressure to penetrate the host cuticle.

Multiple Ehrlichia chaffeensis Genes Critical for Its Persistent Infection in a Vertebrate Host Are Identified by Random Mutagenesis Coupled with In Vivo Infection Assessment.

Ehrlichia chaffeensis, a tick-transmitted obligate intracellular rickettsial agent, causes human monocytic ehrlichiosis. In recent reports, we described substantial advances in developing random and targeted gene disruption methods to investigate the functions of E. chaffeensis genes. We reported earlier that the Himar1 transposon-based random mutagenesis is a valuable tool in defining E. chaffeensis genes critical for its persistent growth in vivo in reservoir and incidental hosts. The method also aided in extending studies focused on vaccine development and immunity. Here, we describe the generation and mapping of 55 new mutations. To define the critical nature of the bacterial genes, infection experiments were carried out in the canine host with pools of mutant organisms. Infection evaluation in the physiologically relevant host by molecular assays and by xenodiagnoses allowed the identification of many proteins critical for the pathogen's persistent in vivo growth. Genes encoding proteins involved in biotin biosynthesis, protein synthesis and fatty acid biosynthesis, DNA repair, electron transfer, and a component of a multidrug resistance (MDR) efflux pump were concluded to be essential for the pathogen's in vivo growth. Three known immunodominant membrane proteins, i.e., two 28-kDa outer membrane proteins (P28/OMP) and a 120-kDa surface protein, were also recognized as necessary for the pathogen's obligate intracellular life cycle. The discovery of many E. chaffeensis proteins crucial for its continuous in vivo growth will serve as a major resource for investigations aimed at defining pathogenesis and developing novel therapeutics for this and related pathogens of the rickettsial family Anaplasmataceae.

An efficient gene disruption method for the woody plant pathogen Botryosphaeria dothidea

BackgroundBotryosphaeria dothidea causes apple white rot and infects many tree plants. Genome data for B. dothidea are available and many pathogenesis-related genes have been predicted. However, a gene manipulation method is needed to study the pathogenic mechanism of B. dothidea.ResultsWe established a gene disruption (GD) method based on gene homologous recombination (GHR) for B. dothidea using polyethylene glycol-mediated protoplast transformation. The results showed that a GHR cassette gave much higher GD efficiency than a GHR plasmid. A high GD efficiency (1.3 ± 0.14 per 106 protopasts) and low frequency of random insertions were achieved with a DNA cassette quantity of 15 μg per 106 protoplasts. Moreover, we successfully disrupted genes in two strains. Bdo_05381-disrupted transformants produced less melanin, whereas the Bdo_02540-disrupted transformant showed a slower growth rate and a stronger resistance to Congo red.ConclusionThe established GD method is efficient and convenient and has potential for studying gene functions and the pathogenic mechanisms of B. dothidea and other coenocytic fungi.

Genome Sequencing of Streptomyces olivaceus SCSIO T05 and Activated Production of Lobophorin CR4 via Metabolic Engineering and Genome Mining.

Marine-sourced actinomycete genus Streptomyces continues to be an important source of new natural products. Here we report the complete genome sequence of deep-sea-derived Streptomyces olivaceus SCSIO T05, harboring 37 putative biosynthetic gene clusters (BGCs). A cryptic BGC for type I polyketides was activated by metabolic engineering methods, enabling the discovery of a known compound, lobophorin CR4 (1). Genome mining yielded a putative lobophorin BGC (lbp) that missed the functional FAD-dependent oxidoreductase to generate the d-kijanose, leading to the production of lobophorin CR4 without the attachment of d-kijanose to C17-OH. Using the gene-disruption method, we confirmed that the lbp BGC accounts for lobophorin biosynthesis. We conclude that metabolic engineering and genome mining provide an effective approach to activate cryptic BGCs.

Generation of deletions and precise point mutations in Dictyostelium discoideum using the CRISPR nickase.

The CRISPR/Cas9 system enables targeted genome modifications across a range of eukaryotes. Although we have reported that transient introduction of all-in-one vectors that express both Cas9 and sgRNAs can efficiently induce multiple gene knockouts in Dictyostelium discoideum, concerns remain about off-target effects and false-positive amplification during mutation detection via PCR. To minimise these effects, we modified the system to permit gene deletions of greater than 1 kb via use of paired sgRNAs and Cas9 nickase. An all-in-one vector expressing the Cas9 nickase and sgRNAs was transiently introduced into D. discoideum, and the resulting mutants showed long deletions with a relatively high efficiency of 10–30%. By further improving the vector, a new dual sgRNA expression vector was also constructed to allow simultaneous insertion of two sgRNAs via one-step cloning. By applying this system, precise point mutations and genomic deletions were generated in the target locus via simultaneous introduction of the vector and a single-stranded oligonucleotide template without integrating a drug resistance cassette. These systems enable simple and straightforward genome editing that requires high specificity, and they can serve as an alternative to the conventional homologous recombination-based gene disruption method in D. discoideum.

Fast gene disruption in Trichoderma reesei using in vitro assembled Cas9/gRNA complex

BackgroundCRISPR/Cas9 has wide application potentials in a variety of biological species including Trichoderma reesei, a filamentous fungus workhorse for cellulase production. However, expression of Cas9 heterologously in the host cell could be time-consuming and sometimes even troublesome.ResultsWe tested two gene disruption methods in T. reesei using CRISPR/Cas9 in this study. The intracellularly expressed Cas9 led to unexpected off-target gene disruption in T. reesei QM9414, favoring inserting 9- or 12-bp at 70- and 100-bp downstream of the targeted ura5. An alternative method was, therefore, established by assembling Cas9 and gRNA in vitro, followed by transformation of the ribonucleoprotein complex with a plasmid containing the pyr4 marker gene into T. reesei TU-6. When the gRNA targeting cbh1 was used, eight among the twenty seven transformants were found to lose the ability to express CBH1, indicative of successful cbh1 disruption through genome editing. Large DNA fragments including the co-transformed plasmid, chromosomal genes, or a mixture of these nucleotides, were inserted in the disrupted cbh1 locus.ConclusionsDirect transformation of Cas9/gRNA complex into the cell is a fast means to disrupt a gene in T. reesei and may find wide applications in strain improvement and functional genomics study.

Role of a membrane-bound aldehyde dehydrogenase complex AldFGH in acetic acid fermentation with Acetobacter pasteurianus SKU1108

Acetic acid fermentation is widely considered a consequence of ethanol oxidation by two membrane-bound enzymes-alcohol dehydrogenase and aldehyde dehydrogenase (ALDH)-of acetic acid bacteria. Here, we used a markerless gene disruption method to construct a mutant of the Acetobacter pasteurianus strain SKU1108 with a deletion in the aldH gene, which encodes the large catalytic subunit of a heterotrimeric ALDH complex (AldFGH), to examine the role of AldFGH in acetic acid fermentation. The ΔaldH strain grew less on ethanol-containing medium, i.e., acetic acid fermentation conditions, than the wild-type strain and significantly accumulated acetaldehyde in the culture medium. Unexpectedly, acetaldehyde oxidase activity levels of the intact ΔaldH cells and the ΔaldH cell membranes were similar to those of the wild-type strain, which might be attributed to an additional ALDH isozyme (AldSLC). The apparent KM values of the wild-type and ΔaldH membranes for acetaldehyde were similar to each other, when the cells were cultured in nonfermentation conditions, where ΔaldH cells grow as well as the wild-type cells. However, the membranes of the wild-type cells grown under fermentation conditions showed a 10-fold lower apparent KM value than those of the cells grown under nonfermentation conditions. Under fermentation conditions, transcriptional levels of a gene for AldSLC were 10-fold lower than those under nonfermentation conditions, whereas aldH transcript levels were not dramatically changed under the two conditions. We suggest that A. pasteurianus SKU1108 has two ALDHs, and the AldFGH complex is indispensable for acetic acid fermentation and is the major enzyme under fermentation conditions.