Microbiology

Macrofungi, also known as mushrooms, can produce various bioactive compounds, including exopolysaccharides (EPS) with distinct biological properties and subsequent industrial applications in the preparation of cosmetics, pharmaceuticals, and food products. EPS are extracellular polymers with diverse chemical compositions and physical properties secreted by macrofungi in the form of capsules or biofilms into the cellular medium. Submerged cultivation is an industrially implemented biotechnological technique used to produce a wide variety of fungal metabolites, which are of economic and social importance due to their food, pharmaceutical, and agronomic applications. It is a favorable technique for cultivating fungi because it requires little space, minimal labor, and low production costs. Moreover, it allows for control over environmental variables and nutrient supply, essential for the growth of the fungus. Although this technique has been widely applied to yeasts, there is limited knowledge regarding optimal growth conditions for filamentous fungi. Filamentous fungi exhibit different behavior compared to yeast, primarily due to differences in cell morphology, reproductive forms, and the type of aggregates generated during submerged fermentation. Furthermore, various growing conditions can affect the production yield of metabolites, necessitating the development of new knowledge to scale up metabolite production from filamentous fungi. This protocol implements the following culture conditions: an inoculum of three agar discs with mycelium, agitation at 150 rpm, a temperature of 28 °C, an incubation time of 72 h, and a carbon source concentration of 40 g/L. These EPS are precipitated using polar solvents such as water, ethanol, and isopropanol and solubilized using water or alkaline solutions. This protocol details the production procedure of EPS using submerged culture; the conditions and culture medium used are described. A detailed description of the extraction is performed, from neutralization to lyophilization. The concentrations and conditions necessary for solubilization are also described.

Key features

• Production and extraction of EPS from submerged cultures of mycelial forms of macrofungi.

• Modification of the method described by Fariña et al. (2001), extending its application to submerged cultures of mycelial forms of the macrofungi.

• Determination of EPS production parameters in submerged cultures of mycelial forms of macrofungi.

• EPS solubilization using NaOH (0.1 N).

Graphical overview

Dolichol diphosphate-linked oligosaccharides (LLO) are the sugar donors in N-glycosylation, a fundamental protein post-translational modification of the eukaryotic secretory pathway. Defects in LLO biosynthesis produce human Congenital Disorders of Glycosylation Type I. The synthesis of LLOs and the transfer reactions to their protein acceptors is highly conserved among animal, plant, and fungi kingdoms, making the fission yeast Schizosaccharomyces pombe a suitable model to study these processes. Here, we present a protocol to determine the LLO patterns produced in vivo by S. pombe cells that may be easily adapted to other cell types. First, exponentially growing cultures are labeled with a pulse of [¹⁴C]-glucose. LLOs are then purified by successive extractions with organic solvents, and glycans are separated from the lipid moieties in mild acid hydrolysis and a new solvent extraction. The purified glycans are then run on paper chromatography. We use a deconvolution process to adjust the profile obtained to the minimal number of Gaussian functions needed to fit the data and determine the proportion of each species with respect to total glycan species present in the cell. The method we provide here might be used without any expensive or specialized equipment. The deconvolution process described here might also be useful to analyze species in non-completely resolved chromatograms.

Graphical abstract:

Workflow for the labeling, extraction, separation, and identification of LLO species in S. pombe. (A) Radioactive pulse of S. pombe cells with [¹⁴C]-glucose for 15 min at 28 °C. (B) Organic extraction of LLOs from labeled yeasts sequentially using methanol, chloroform, H₂O, chloroform:methanol:H₂O (1:1:0.3), 0.02 M HCl (to separate glycans from dolichol), and chloroform:methanol:H₂O (1:16:16). (C) Preparation of the sample for chromatography on paper: drying by airflow and radioactivity check. (D) Loading of samples in chromatographic paper and descendent chromatography in a glass chamber. The obtained plots (CPM versus running distance) need to be analyzed to identify single glycan species.

Lipopolysaccharides (LPS) (or lipooligosaccharides [LOS], which lack the O-antigen side chains characteristic of LPS), and outer membrane proteins (OMP) are major cell-surface molecules in the outer membrane (OM) of gram-negative bacteria. The LPS is responsible for causing endotoxic shock in infected hosts and, in conjunction with some OMPs, provides protection to the bacterium against host innate immune defenses and attachment to host cells. Electrophoretic analysis can provide valuable information regarding the size, number, and variability of LPS/LOS and OMP components between bacterial strains and mutants, which aids in understanding the basic biology and virulence factors of a particular species. Furthermore, highly purified extracts are normally not required if only electrophoretic analysis is to be done, and various methods have been established for such procedures. Here, we review ameliorated procedures for fast and convenient extraction of LPS/LOS and protein-enriched outer membranes (PEOM) for optimal electrophoretic resolution. Specifically, we will describe the phenol-based micro-method for LPS/LOS extraction, a differential extraction procedure with sodium lauryl sarcosinate for PEOM, and gel preparation for electrophoretic analysis of LPS/LOS samples in detail.

Graphic abstract:

Workflow for the preparation and analysis of LPS/LOS and PEOM.

This is a protocol for quantitative determination of storage and total carbohydrates in algae and cyanobacteria. The protocol is simple, fast and sensitive and it requires only few standard chemicals. Great advantage of this protocol is that both storage and total saccharides can be determined in the cellular pellets that were already used for chlorophyll and carotenoids quantification. Since it is recommended to perform the pigments measurement in triplicates, each pigment analysis can generate samples for both total saccharide and glycogen/starch content quantification.

The protocol was applied for quantification of both storage and total carbohydrates in cyanobacteria Synechocystis sp. PCC 6803, Cyanothece sp. ATCC 51142 and Cyanobacterium sp. IPPAS B-1200. It was also applied for estimation of storage polysaccharides in Galdieria (IPPAS P-500, IPPAS P-507, IPPAS P-508, IPPAS P-513), Cyanidium caldarium IPPAS P-510, in green algae Chlorella sp. IPPAS C-1 and C-1210, Parachlorella kessleri IPPAS C-9, Nannochloris sp. C-1509, Coelastrella sp. IPPAS H-626, Haematococcus sp. IPPAS H-629 and H-239, and in Eustigmatos sp. IPPAS H-242 and IPPAS C-70.

Invertase can catalyze the hydrolysis of sucrose, and is widely distributed in cells of cyanobacteria and plants. Being responsible for the first step for sucrose metabolism, invertase plays important physiological roles and its enzymatic activity is frequently needed to be determined. All the methods for determination of the invertase activity are dependent on detection of the glucose product generated by the invertase. Here we describe an ion chromatography based protocol of our laboratory for determination of cyanobacterial intracellular invertase activity.

Cyanobacteria, which have the extraordinary ability to grow using sunlight and carbon dioxide, are emerging as a green host to produce value-added products. Exploitation of this highly promising host to make products may depend on the ability to modulate the glucose metabolic pathway; it is the key metabolic pathway that generates intermediates that feed many industrially important pathways. Thus, before cyanobacteria can be considered as a leading source to produce value-added products, we must understand the interaction between glucose metabolism and other important cellular activities such as photosynthesis and chlorophyll metabolism. Here we describe reproducible and reliable methods for measuring extracellular glucose and glycogen levels from cyanobacteria.

The marine beta-glucan laminarin is an abundant storage polysaccharide in microalgae. High production rates and rapid digestion by heterotrophic bacteria turn laminarin into an ideal carbon and energy source, and it is therefore a key player in the marine carbon cycle. As a main storage glucan laminarin also plays a central role in the energy metabolism of the microalgae (Percival and Ross, 1951; Myklestad, 1974; Painter, 1983). We take advantage of enzymes that digest laminarin selectively and can thereby quantify only this polysaccharide in environmental samples. These enzymes hydrolyze laminarin into glucose and oligosaccharides, which are measured with a standard reducing sugar assay to obtain the laminarin concentration. Prior to this assay, the three enzymes need to be produced via heterologous expression and purification. The assay can be used to monitor laminarin concentrations in environmental microalgae, which were concentrated from seawater by filtering, or in samples derived from algal lab cultures.

Peptidoglycan encases the bacterial cytoplasmic membrane to protect the cell from lysis due to the turgor. The final steps of peptidoglycan synthesis require a membrane-anchored substrate called lipid II, in which the peptidoglycan subunit is linked to the carrier lipid undecaprenol via a pyrophosphate moiety. Lipid II is the target of glycopeptide antibiotics and several antimicrobial peptides, and is degraded by ‘attacking’ enzymes involved in bacterial competition to induce lysis. Here we describe two protocols using thin-layer chromatography (TLC) and high pressure liquid chromatography (HPLC), respectively, to assay the digestion of lipid II by phosphatases such as Colicin M or the LXG toxin protein TelC from Streptococcus intermedius. The TLC method can also monitor the digestion of undecaprenyl (pyro)phosphate, whereas the HPLC method allows to separate the di-, mono- or unphosphorylated disaccharide pentapeptide products of lipid II.

Streptococcus pneumoniae (pneumococcus) is an important human pathogen that causes pneumonia, meningitis, sepsis, and otitis media. This bacterium normally resides in the nasopharynx as a commensal, but sometimes disseminates to sterile sites of humans and causes local or systemic inflammation. This biphasic behavior of S. pneumoniae is correlated with a reversible switch between the opaque and transparent colony forms on agar plates, a phenomenon referred to as phase variation. The opaque variants appear to be more virulent in animal models of bacteremia but are deficient in nasopharyngeal colonization animal models. In contrast, the transparent variants display higher levels of nasopharyngeal colonization but relatively lower virulence in animal models. We have recently demonstrated that pneumococcal phase variation between these two colony types is caused by a reversible switch of genome DNA methylation (or epigenetic) patterns, which is driven by DNA inversions in the DNA methyltransferase genes. Observation of colony morphology is a simple and useful method to differentiate colonies with different characteristics, such as size, color, and opacity. This protocol describes how to study pneumococcal phase variation in colony morphology with a dissection microscope.

Peptidoglycan (murein) is a vital component of the cell wall of nearly all bacteria, composed of sugars linked by short peptides. This protocol describes the purification of macromolecular peptidoglycan from cultured bacteria and the analysis of enzyme-digested peptidoglycan fragments using high performance liquid chromatography (HPLC). Digested peptidoglycan fragments can be identified by mass spectrometry, or predicted by comparing retention times with other published chromatograms. The quantitative nature of this method allows for the measurement of changes to peptidoglycan composition between different species of bacteria, growth conditions, or mutations. This method can determine the overall architecture of peptidoglycan, such as peptide stem length, the extent of cross-linking, and modifications. Muropeptide analysis has been used to study the function of peptidoglycan-associated proteins and the mechanisms by which bacteria acquire antibiotic resistance.