Biochemistry

Candida glabrata is an opportunistic pathogen that may cause serious infections in an immunocompromised host. C. glabrata cell wall proteases directly interact with host cells and affect yeast virulence and host immune responses. This protocol describes methods to purify β-1,3-glucan-bonded cell wall proteases from C. glabrata. These cell wall proteases are detached from the cell wall glucan network by lyticase treatment, which hydrolyzes β-1,3-glucan bonds specifically without rupturing cells. The cell wall supernatant is further fractioned by centrifugal devices with cut-offs of 10 and 50 kDa, ion-exchange filtration(charge), and gel filtration (size exclusion). The enzymatic activity of C. glabrata proteases is verified with MDPF-gelatin zymography and the degradation of gelatin is visualized by loss of gelatin fluorescence. With this procedure, the enzymatic activities of the fractions are kept intact, differing from methods used in previous studies with trypsin digestion of the yeast cell wall. The protein bands may be eventually located from a parallel silver-stained gel and identified with LC–MS/MS spectrometry. The advantage of this methodology is that it allows further host protein degradation assays; the protocol is also suitable for studying other Candida yeast species.

Key features

• Uses basic materials and laboratory equipment, enabling low-cost studies.

• Facilitates the selection and identification of proteases with certain molecular weights.

• Enables further functional studies with host proteins, such as structural or immune response–related, or enzymes and candidate protease inhibitors(e.g., from natural substances).

• This protocol has been optimized for C. glabrata but may be applied with modifications to other Candida species.

Graphical overview

Nanobodies are recombinant antigen-specific single domain antibodies (VHHs) derived from the heavy chain–only subset of camelid immunoglobulins. Their small molecular size, facile expression, high affinity, and stability have combined to make them unique targeting reagents with numerous applications in the biomedical sciences. From our work in producing nanobodies to over sixty different proteins, we present a standardised workflow for nanobody discovery from llama immunisation, library building, panning, and small-scale expression for prioritisation of binding clones. In addition, we introduce our suites of mammalian and bacterial vectors, which can be used to functionalise selected nanobodies for various applications such as in imaging and purification.

Key features

• Standardise the process of building nanobody libraries and finding nanobody binders so that it can be repeated in any lab with reasonable equipment.

• Introduce two suites of vectors to functionalise nanobodies for production in either bacterial or mammalian cells.

Graphical overview

Enzyme immobilization offers a number of advantages that improve biocatalysis; however, finding a proper way to immobilize enzymes is often a challenging task. Implanting enzymes in metal–organic frameworks (MOFs) via co-crystallization, also known as biomineralization, provides enhanced reusability and stability with minimal perturbation and substrate selectivity to the enzyme. Currently, there are limited metal–ligand combinations with a proper protocol guiding the experimental procedures. We have recently explored 10 combinations that allow custom immobilization of enzymes according to enzyme stability and activity in different metals/ligands. Here, as a follow-up of that work, we present a protocol for how to carry out custom immobilization of enzymes using the available combinations of metal ions and ligands. Detailed procedures to prepare metal ions, ligands, and enzymes for their co-crystallization, together with characterization and assessment, are discussed. Precautions for each experimental step and result analysis are highlighted as well. This protocol is important for enzyme immobilization in various research and industrial fields.

Key features

• A wide selection of metal ions and ligands allows for the immobilization of enzymes in metal–organic frameworks (MOFs) via co-crystallization.

• Step-by-step enzyme immobilization procedure via co-crystallization of metal ions, organic linkers, and enzymes.

• Practical considerations and experimental conditions to synthesize the enzyme@MOF biocomposites are discussed.

• The demonstrated method can be generalized to immobilize other enzymes and find other metal ion/ligand combinations to form MOFs in water and host enzymes.

Graphical overview

The human pathogenic yeast Candida albicans can attach to epithelial cells or indwelling medical devices to form biofilms. These microbial communities are highly problematic in the clinic as they reduce both sensitivity to antifungal drugs and detection of fungi by the immune system. Amyloid structures are highly organized quaternary structures that play a critical role in biofilm establishment by allowing fungal cells to adhere to each other. Thus, fungal amyloids are exciting targets to develop new antifungal strategies. Thioflavin T is a specific fluorescent dye widely used to study amyloid properties of target proteins in vitro (spectrophotometry) and in vivo (epifluorescence/confocal microscopy). Notably, thioflavin T has been used to demonstrate the ability of Als5, a C. albicans adhesin, to form an amyloid fiber upon adhesion. We have developed a pipeline that allows us to study amyloid properties of target proteins using thioflavin T staining in vitro and in vivo, as well as in intact fungal biofilms. In brief, we used thioflavin T to sequentially stain (i) amyloid peptides, (ii) recombinant proteins, (iii) fungal cells treated or not with amyloid peptides, (iv) fungal amyloids enriched by cell fractionation, and (v) intact biofilms of C. albicans. Contrary to other methods, our pipeline gives a complete picture of the amyloid behavior of target proteins, from in vitro analysis to intact fungal biofilms. Using this pipeline will allow an assessment of the relevance of the in vitro results in cells and the impact of amyloids on the development and/or maintenance of fungal biofilm.

Key features

• Study of amyloid properties of fungal proteins.

• Visualization of the subcellular localization of fungal amyloid material using epifluorescence or confocal microscopy.

• Unraveling of the amyloid properties of target proteins and their physiological meaning for biofilm formation.

• Observation of the presence of amyloid structures with live-cell imaging on intact fungal biofilm using confocal microscopy.

Graphical overview

As the most energy- and metabolite-consuming process, protein synthesis is under the control of several intrinsic and extrinsic factors that determine its fine-tuning to the cellular microenvironment. Consequently, variations in protein synthesis rates occur under various physiological and pathological conditions, enabling an adaptive response by the ce•ll. For example, global protein synthesis increases upon mitogenic factors to support biomass generation and cell proliferation, while exposure to low concentrations of oxygen or nutrients require translational repression and reprogramming to avoid energy depletion and cell death. To assess fluctuations in protein synthesis rates, radioactive isotopes or radiolabeled amino acids are often used. Although highly sensitive, these techniques involve the use of potentially toxic radioactive compounds and require specific materials and processes for the use and disposal of these molecules. The development of alternative, non-radioactive methods that can be easily and safely implemented in laboratories has therefore been encouraged to avoid handling radioactivity. In this context, the SUrface SEnsing of Translation (SUnSET) method, based on the classical western blot technique, was developed by Schmidt et al. in 2009. The SUnSET is nowadays recognized as a simple alternative to radioactive methods assessing protein synthesis rates.

Key features

• As a structural analogue of aminoacyl-transfer RNA, puromycin incorporates into the elongating peptide chain.

• Detection of puromycin-labeled peptides by western blotting reflects translation rates without the need for radioactive isotopes.

• The protocol described here for in vitro applications is derived from the SUnSET method originally published by Schmidt et al. (2009).

Proteolysis is a critical biochemical process yet a challenging field to study experimentally due to the self-degradation of a protease and the complex, dynamic degradation steps of a substrate. Mass spectrometry (MS) is the traditional way for proteolytic studies, yet it is challenging when time-resolved, step-by-step details of the degradation process are needed. We recently found a way to resolve the cleavage site, preference/selectivity of cleavage regions, and proteolytic kinetics by combining site-directed spin labeling (SDSL) of protein substrate, time-resolved two-dimensional (2D) electron paramagnetic resonance (EPR) spectroscopy, protease immobilization via metal–organic materials (MOMs), and MS. The method has been demonstrated on a model substrate and protease, yet there is a lack of details on the practical operations to carry out our strategy. Thus, this protocol summarizes the key steps and considerations when carrying out the EPR/MS study on proteolytic processes, which can be generalized to study other protein/polypeptide substrates in proteolysis. Details for the experimental operation and cautions of each step are reported with figures illustrating the concepts. This protocol provides an effective approach to understanding the proteolytic process with the advantages of offering time-resolved, residue-level resolution of structural basis underlying the process. Such information is important for revealing the cleavage site and proteolytic mechanisms of unknown proteases. The advantage of EPR, probing the target substrate regardless of the complexities caused by the proteases and their self-degradation, offers a practically effective, rapid, and easy-to-operate approach to studying proteolysis.

Key features

• Combining protease immobilization, EPR, spin labeling, and MS experimental methods allows for the analysis of proteolysis process in real time.

• Reveals cleavage site, kinetics of product generation, and preference of cleavage regions via time-resolved SDSL-EPR.

• MS confirms EPR findings and helps depict the sequences and populations of the cleaved segments in real time.

• The demonstrated method can be generalized to other proteins or polypeptide substrates upon proteolysis by other proteases.

Graphical overview

Tears contain numerous secreted factors, enzymes, and proteins that help in maintaining the homeostatic condition of the eye and also protect it from the external environment. However, alterations to these enzymes and/or proteins during pathologies such as mechanical injury and viral or fungal infections can disrupt the normal ocular homeostasis, further contributing to disease development. Several tear film components have a significant role in curbing disease progression and promoting corneal regeneration. Additionally, several factors related to disease progression are secreted into the tear film, thereby serving as a valuable reservoir of biomarkers. Tears are readily available and can be collected via non-invasive techniques or simply from contact lenses. Tears can thus serve as a valuable and easy source for studying disease-specific biomarkers. Significant advancements have been made in recent years in the field of tear film proteomics, lipidomics, and transcriptomics to allow a better understanding of how tears can be utilized to gain insight into the etiology of diseases. These advancements have enabled us to study the pathophysiology of various disease states using tear samples. However, the mechanisms by which tears help to maintain corneal homeostasis and how they are able to form the first line of defense against pathogens remain poorly understood and warrant detailed in vitro studies. Herein, we have developed an in vitro assay to characterize the functional importance of patient isolated tears and their components on corneal epithelial cells. This novel approach closely mimics real physiological conditions and could help the researchers gain insight into the underlying mechanisms of ocular pathologies and develop new treatments.

Key features

• This method provides a new technique for analyzing the effect of tear components on human corneal epithelial cells.

• The components of the tears that are altered in response to diseases can be used as a biomarker for detecting ocular complications.

• This procedure can be further employed as an in vitro model for assessing the efficacy of drugs and discover potential therapeutic interventions.

Autophagy is an essential catabolic pathway used to sequester and engulf cytosolic substrates via a unique double-membrane structure, called an autophagosome. The ubiquitin-like ATG8 proteins play an important role in mediating autophagosome membrane expansion. They are covalently conjugated to phosphatidylethanolamine (PE) on the autophagosomes via a ubiquitin-like conjugation system called ATG8 lipidation. In vitro reconstitution of ATG8 lipidation with synthetic liposomes has been previously established and used widely to characterise the function of the E1 ATG7, the E2 ATG3, and the E3 complex ATG12–ATG5-ATG16L1. However, there is still a lack of a tool to provide kinetic measurements of this enzymatic reaction. In this protocol, we describe a real-time lipidation assay using NBD-labelled ATG8. This real-time assay can distinguish the formation of ATG8 intermediates (ATG7~ATG8 and/or ATG3~ATG8) and, finally, ATG8-PE conjugation. It allows kinetic characterisation of the activity of ATG7, ATG3, and the E3 complex during ATG8 lipidation. Furthermore, this protocol can be adapted to characterise the upstream regulators that may affect protein activity in ATG8 lipidation reaction with a kinetic readout.

Key features

• Preparation of ATG7 E1 from insect cells (Sf9 cells).

• Preparation of ATG3 E2 from bacteria (E. coli).

• Preparation of LC3B S3C from bacteria (E. coli).

• Preparation of liposomes to monitor the kinetics of ATG8 lipidation in a real-time manner.

Graphical overview

Experimental design to track the full reaction of ATG8 lipidation, described in this protocol

In situ cryo-electron tomography (cryo-ET) is the most current, state-of-the-art technique to study cell machinery in its hydrated near-native state. The method provides ultrastructural details at sub-nanometer resolution for many components within the cellular context. Making use of recent advances in sample preparation techniques and combining this method with correlative light and electron microscopy (CLEM) approaches have enabled targeted molecular visualization. Nevertheless, the implementation has also added to the complexity of the workflow and introduced new obstacles in the way of streamlining and achieving high throughput, sample yield, and sample quality. Here, we report a detailed protocol by combining multiple newly available technologies to establish an integrated, high-throughput, optimized, and streamlined cryo-CLEM workflow for improved sample yield.

Key features

• PRIMO micropatterning allows precise cell positioning and maximum number of cell targets amenable to thinning with cryo focused-ion-beam–scanning electron microscopy.

• CERES ice shield ensures that the lamellae remain free of ice contamination during the batch milling process.

• METEOR in-chamber fluorescence microscope facilitates the targeted cryo focused-ion-beam (cryo FIB) milling of these targets.

• Combining the three technologies into one cryo-CLEM workflow maximizes sample yield, throughput, and efficiency.

Graphical overview

Advanced immunoassays are crucial in assessing antibody responses, serving immune surveillance goals, characterising immunological responses to evolving viral variants, and guiding subsequent vaccination initiatives. This protocol outlines an indirect ELISA protocol to detect and quantify virus-specific antibodies in plasma or serum after exposure to viral antigens. The assay enables the measurement of IgG, IgA, and IgM antibodies specific to the virus of interest, providing qualitative and quantitative optical densities and concentration data. Although this protocol refers to SARS-CoV-2, its methodology is versatile and can be modified to assess antibody responses for various viral infections and to evaluate vaccine trial outcomes.

Key features

• This protocol builds upon previously described methodology [1] explicitly tailored for SARS-CoV-2 and broadens its applicability to other viral infections.

• The protocol outlines establishing antibody responses to SARS-CoV-2 infections by determining optical densities and concentrations from blood plasma or serum.

Graphical overview

Summary of the conventional ELISA (A) vs. sensitive ELISA (B) procedures. In both A and B, wells are coated with a capture antigen, such as the spike protein, while in (C) they are coated with human Kappa and Lambda capture antibodies. For the conventional ELISA (A), wells with immobilised capture antigens receive serum/plasma containing the target antibody (A1 and B1). This is followed by an HRP-conjugated detection antibody specific to the captured antibody (A2 and B2) and then a substrate solution that reacts with the HRP, producing a colour proportional to the concentration of the antibody in the serum/plasma (A3 and B3). The reaction is halted, and absorbance is measured. In the sensitive ELISA (B), after the serum/plasma addition (A1 and B1), a Biotin-conjugated primary detection antibody is introduced (A2 and B2). Depending on the target antibody, a secondary streptavidin-HRP conjugated detection antibody is added for IgG or IgM (3a) or a poly-HRP 40 detection antibody for IgA (3b). A substrate is introduced, producing a colour change proportional to the antibody concentration (A4 and B4). The reaction is then stopped, and absorbance is measured. In Panel C, wells are coated with human Kappa and Lambda capture antibodies. Serial dilutions of a known antibody standard are introduced. After undergoing the standard ELISA steps, a detection antibody is added, specifically binding to the Ig standard antibody. Subsequently, a substrate solution causes a colour change proportional to the antibody concentration in the serum/plasma. The reaction is halted, and the absorbance of each well is measured. The resulting optical densities from the coated wells form the standard curve, plotting the absorbance against concentrations.