Biochemistry

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.

Far-western blotting, derived from the western blot, has been used to detect interactions between proteins in vitro, such as receptor–ligand interactions. The insulin signaling pathway plays a critical role in the regulation of both metabolism and cell growth. The binding of the insulin receptor substrate (IRS) to the insulin receptor is essential for the propagation of downstream signaling after the activation of the insulin receptor by insulin. Here, we describe a step-by-step far-western blotting protocol for determining the binding of IRS to the insulin receptor.

The subretinal layer between retinal pigment epithelium (RPE) and photoreceptors is a region involved in inflammation and angiogenesis during the procession of diseases such as age-related macular degeneration. The current protocols of whole mounts (retina and RPE) are unable to address the intact view of the subretinal layer because the separation between retina and RPE is required, and each separate tissue is then stained. Non-separate Sclerochoroid/RPE/Retina whole mount staining was recently developed and reported. The method can be further combined and optimized with melanin bleaching and tissue clearing. Here, we describe steps of both non-pigmented and pigmented mouse Sclerochoroid/RPE/Retina whole mount including eyeball preparation, staining, mounting and confocal scanning. In addition, we present the confocal images of RPE, subretinal microglia and the neighboring photoreceptors in Sclerochoroid/RPE/Retina whole mounts.

Protein-protein interactions constitute the molecular foundations of virtually all biological processes. Co-immunoprecipitation (CoIP) experiments are probably the most widely used method to probe both heterotypic and homotypic protein-protein interactions. Recent advances in super-resolution microscopy have revealed that several nuclear proteins such as transcription factors are spatially distributed into local high-concentration clusters in mammalian cells, suggesting that many nuclear proteins self-interact. These observations have further underscored the need for orthogonal biochemical approaches for testing if self-association occurs, and if so, what the mechanisms are. Here, we describe a CoIP protocol specifically optimized to test self-association of endogenously tagged nuclear proteins (self-CoIP), and to evaluate the role of nucleic acids in such self-interaction. This protocol has proven reliable and robust in our hands, and it can be used to test both homotypic and heterotypic (CoIP) protein-protein interactions.

Mammalian skeletal muscle is a metabolically active tissue that is made up of different types of muscle fibers. These myofibers are made up of important contractile proteins that provide force during contraction of the muscle like actin and myosin. Murine myofibers have been classified into 4 types: Type I, Type IIa, Type IIb and Type IIX. Each muscle fiber has been identified with specific type of MyHC expressed, which in turn gives differential contractility to the muscle.

There have been well-known methodologies to identify different myofibers: histochemical myosin ATPase staining which uses the differential ATPase activity between slow and fast fibers, quantification of metabolic enzymes like malate dehydrogenase and lactate dehydrogenase on specific fragments of muscle fibers. The drawback of these techniques is that they cannot differentiate the subtypes of myofibers, for example, Type IIa and Type IIb. They should be used in conjunction with other known histochemical staining techniques. Here, we devise a direct and robust immunohistochemical staining methodology that utilizes the differential expression of MyHC isoforms in different myofibers types, thus efficiently distinguishing the heterogeneity of the muscle fibers. We use antibodies that specifically recognize Type I, Type IIa and Type IIb fibers on serially cut frozen mouse tibialis anterior sections that can be quantified by ImageJ software.

Oxidative stress is associated with numerous diseases, and markers of oxidative stress in biological material are becoming a mainstay of both experimental and clinical/epidemiological research. Lipid peroxidation is a major form of oxidative stress, but due to their rapid degradation and instability, lipid peroxides are notoriously difficult to measure, particularly in biological specimens where their production and removal are continuously occuring. Thus, a commonly used surrogate marker of lipid peroxidation is protein adducts of 4-Hydroxynonenal (HNE), an α, β-unsaturated hydroxyalkenal (i.e., a reactive aldehyde) formed via degradation of oxidized polyunsaturated fatty acids (PUFAs). HNE adducts can be measured via commercially-available immunosorbent assays, but these have their limitations due to excessive costs, and reproducibility among laboratories is challenging due to variability in assay sensitivity, procedure, and reagents. Here we present a reproducible, facile, and economically conservative protocol for quantifying HNE protein adducts. The key to this protocol is to generate HNE-adduct standards by incubating bovine serum albumin (BSA) with HNE. These standards are then adsorbed to immunsorbent plastic in a multi-well plate format alongside biological samples. An enzyme-linked immunosorbent assay (ELISA) is then performed on the multi-well plate using commercially-available primary and secondary antibodies, and a peroxide-based fluorescent developing reagent. This protocol is highly sensitive and offers advantages to commercial sources in that it allows for reproducible, high-throughput quantitation of HNE adducts in a large number of samples.

Protein analysis in bone marrow samples from patients with multiple myeloma (MM) has been limited by the low concentration of proteins obtained after CD138⁺ cell selection. A novel approach based on capillary nano-immunoassay could make it possible to quantify dozens of proteins from each CD138⁺ purified MM sample in an automated manner. Up to now, the knowledge of protein level in those cells was limited because a relatively small quantity of sample is available after the diagnostic procedure. Moreover, the sample often is required for nucleic acids analysis. We have developed the procedure for obtaining proteins from bone marrow samples preserved in RLT+ buffer, and we have successfully applied this approach for the quantification of proteins in the setting of patients with MM. Proteins are extracted from RLT+ buffer, the content is quantified by total protein assay with WES machine and finally, the particular protein expression level is evaluated using specific antibodies by capillary nano-immunoassay with WES machine. The present protocol enables us to quantify many proteins from a limited amount of sample, without losing the opportunity to obtain nucleic acids at the same time. Proteins are quantified automatically in an assay with a low probability of human errors, which makes it a useful tool for biomarkers development.

The recent discovery of human signal peptide peptidase-like 2a (SPPL2a) deficiency in humans revealed the toxicity associated with the accumulation of one of its substrates, CD74 N-terminal fragment (CD74-NTF), for certain type of dendritic cells (cDC2). We developed a two-step protocol for monitoring the accumulation of this molecule in different subsets of PBMCs and immortalized B cells, in which SPPL2a is chemically inhibited and CD74-NTF levels are then assessed by flow cytometry or western blotting. The chemical inhibition of SPPL2a has been described elsewhere, but this is the first time that this inhibition has been reported as a protocol.

Aquatic organisms have specialized cells called ionocytes that regulate the ionic composition, osmolarity, and acid/base status of internal fluids. In small aquatic organisms such as fishes in their early life stages, ionocytes are typically found on the cutaneous surface and their abundance can change to help cope with various metabolic and environmental factors. Ionocytes profusely express ATPase enzymes, most notably Na⁺/K⁺ ATPase, which can be identified by immunohistochemistry. However, quantification of cutaneous ionocytes is not trivial due to the limited camera’s focal plane and the microscope’s field-of-view. This protocol describes a technique to consistently and reliably identify, image, and measure the relative surface area covered by cutaneous ionocytes through software-mediated focus-stacking and photo-stitching–thereby allowing the quantification of cutaneous ionocyte area as a proxy for ion transporting capacity across the skin. Because ionocytes are essential for regulating ionic composition, osmolarity, and acid/base status of internal fluids, this technique is useful for studying physiological mechanisms used by fish larvae and other small aquatic organisms during development and in response to environmental stress.

Biofilm formation is a well-known bacterial strategy that protects cells from hostile environments. During infection, bacteria found in a biofilm community are less sensitive to antibiotics and to the immune response, often allowing them to colonize and persist in the host niche. Not surprisingly, biofilm formation on medical devices, such as urinary catheters, is a major problem in hospital settings. To be able to eliminate such biofilms, it is important to understand the key bacterial factors that contribute to their formation. A common practice in the lab setting is to study biofilms grown in laboratory media. However, these media do not fully reflect the host environment conditions, potentially masking relevant biological determinants. This is the case during urinary catheterization, where a key element for Enterococcus faecalis and Staphylococcus aureus colonization and biofilm formation is the release of fibrinogen (Fg) into the bladder and its deposition on the urinary catheter. To recapitulate bladder conditions during catheter-associated urinary tract infection (CAUTI), we have developed a fibrinogen-coated catheter and 96-well plate biofilm assay in urine. Notably, enterococcal biofilm factors identified in these in vitro assays proved to be important for biofilm formation in vivo in a mouse model of CAUTI. Thus, the method described herein can be used to uncover biofilm-promoting factors that are uniquely relevant in the host environment, and that can be exploited to develop new antibacterial therapies.