Premed · Premed · Immunology
Lecture 30: Immunological Techniques: ELISA, Flow Cytometry, Western Blot
Immunology
Learning Objectives
By the end of this lecture, students will be able to:
- Explain the principle and types of ELISA (direct, indirect, sandwich, competitive) and their clinical applications
- Describe the principles of flow cytometry and its use in immunophenotyping
- Explain the Western blot technique and its application as a confirmatory diagnostic test
- Describe additional immunological techniques including immunofluorescence, immunohistochemistry, and immunoprecipitation
- Interpret results from common immunological assays in clinical and research settings
Lecture Content
I. Principles Common to Immunological Techniques
Most immunological techniques exploit the specificity of antibody-antigen binding, using antibodies as tools or reagents to detect, quantify, or localize target molecules. Several key concepts underpin these methods. A primary antibody binds the target antigen directly. A secondary antibody binds the primary antibody (for example, a goat anti-mouse IgG) and serves to amplify the signal. Antibodies are linked to detectable labels through conjugation, with common labels including enzymes (HRP, horseradish peroxidase; AP, alkaline phosphatase) for colorimetric or chemiluminescent detection, fluorophores (FITC, PE, APC, Alexa Fluor dyes) for fluorescence detection, and biotin (detected by streptavidin conjugates) for signal amplification.
Two types of antibody reagents are available. Monoclonal antibodies, produced from a single B cell clone using hybridoma technology (developed by Kohler and Milstein, Nobel Prize 1984), recognize a single epitope and provide high specificity. Polyclonal antibodies are mixtures from multiple B cell clones that recognize multiple epitopes, offering higher sensitivity but less specificity.
II. Enzyme-Linked Immunosorbent Assay (ELISA)
ELISA is a quantitative assay that uses enzyme-linked antibodies to detect and measure antigens or antibodies in a sample. It is performed in 96-well microtiter plates and exists in four main formats.
Direct ELISA is the simplest format. Antigen is coated on the plate, and an enzyme-conjugated primary antibody is added that binds the antigen directly. When substrate is added, the resulting color change is proportional to the amount of antigen present. While simple, this format is less sensitive and less commonly used in practice.
Indirect ELISA is widely used in clinical diagnostics. Antigen is coated on the plate, then patient serum containing unknown antibodies is added. If antibodies specific for the coated antigen are present, they bind. An enzyme-conjugated secondary antibody (such as anti-human IgG-HRP) is then added to detect the patient's bound antibodies. Color change upon substrate addition indicates the presence of antibody in the patient serum. This format is used clinically to detect antibodies against pathogens, including anti-HIV antibodies, anti-HCV, and anti-viral IgG/IgM. Its advantages include versatility, since the same secondary antibody works for any primary, and inherent signal amplification.
Sandwich ELISA is the most sensitive and specific format. A capture antibody is coated on the plate that binds one epitope on the target antigen. The sample containing the antigen is added and captured. A detection antibody conjugated to an enzyme is then added that binds a different epitope on the antigen, creating a "sandwich." The color change produced by the substrate is proportional to the antigen concentration. This format is used clinically to quantify cytokines (IL-6, TNF-alpha), hormones, and tumor markers (PSA, CEA).
Competitive ELISA uses a pre-coated plate with antigen. Patient sample and enzyme-conjugated antibody are added simultaneously. If antigen is present in the patient sample, it competes with the coated antigen for antibody binding, resulting in less antibody bound to the plate and therefore less color development. The signal is inversely proportional to the antigen concentration in the sample. This format is particularly useful when the antigen is small (a hapten) or when only one epitope is available for binding.
Results are read using a spectrophotometer (plate reader) that measures optical density, with quantification achieved through comparison to a standard curve. ELISA sensitivity typically reaches the pg/mL to ng/mL range, which can be enhanced further by using chemiluminescent substrates.
<image>A four-panel diagram comparing the four types of ELISA. Panel A (Direct ELISA): A microtiter well is shown with antigen molecules (triangles) coated on the bottom. An enzyme-conjugated primary antibody (Y-shape with enzyme star) binds directly to the antigen. Substrate is added, producing a colored product. Panel B (Indirect ELISA): Antigen is coated on the well. Patient serum antibodies (red Y-shapes) bind the antigen. An enzyme-conjugated secondary antibody (green Y-shape with enzyme star, labeled "anti-human IgG-HRP") binds the patient antibody. Substrate produces color. Panel C (Sandwich ELISA): A capture antibody (blue Y-shape) is coated on the well. Antigen from the sample (triangle) binds the capture antibody. A different enzyme-conjugated detection antibody (green Y-shape with enzyme star) binds a second epitope on the antigen, creating a sandwich. Substrate produces color proportional to antigen concentration. A standard curve graph is shown alongside for quantification. Panel D (Competitive ELISA): Antigen is coated on the well. A mixture of sample (containing free antigen) and enzyme-conjugated antibody is added. Free antigen in the sample competes with coated antigen for antibody binding. Less antibody binds the plate when more free antigen is present. Inverse relationship graph is shown: higher sample antigen → less color.</image>
III. Flow Cytometry
Flow cytometry is a technique in which individual cells in suspension pass single-file through a laser beam, and scattered light and fluorescence emissions are measured for each cell, enabling multiparameter analysis of thousands of cells per second. Sample preparation involves creating a single-cell suspension from blood, bone marrow, tissue digest, or cultured cells, which is then stained with fluorophore-conjugated antibodies against surface or intracellular markers.
A flow cytometer consists of three main systems. The fluidics system uses hydrodynamic focusing to create a single-file stream of cells. The optics system includes lasers that excite fluorophores and detectors that measure the resulting signals. Forward scatter (FSC) is proportional to cell size, while side scatter (SSC) is proportional to cell granularity or complexity. Multiple fluorescence detectors capture emission from fluorophore-conjugated antibodies across different color channels. The electronics and software convert signals to digital data and display them as dot plots, histograms, or contour plots.
Immunophenotyping is the most common clinical application of flow cytometry. It identifies and quantifies cell populations based on surface marker expression. The CD4+ T cell count is essential for monitoring HIV/AIDS progression, with AIDS defined as CD4 below 200 per microliter. Leukemia and lymphoma immunophenotyping determines cell lineage (B cell, T cell, myeloid), maturation stage, and aberrant markers to classify hematologic malignancies. Lymphocyte subset analysis identifies CD3+ T cells, CD4+ helper T cells, CD8+ CTLs, CD19+ B cells, and CD16+CD56+ NK cells. Stem cell enumeration of CD34+ cells is performed for HSCT. Paroxysmal nocturnal hemoglobinuria (PNH) is diagnosed by detecting loss of GPI-anchored proteins (CD55, CD59) on red blood cells and white blood cells.
Advanced applications of flow cytometry include intracellular cytokine staining, in which cells are fixed and permeabilized to stain for intracellular cytokines (IFN-gamma, IL-4, IL-17), identifying T cell subsets by function. FOXP3 staining identifies Tregs through intracellular staining after permeabilization. Proliferation assays use CFSE dye dilution, where each cell division halves the fluorescence intensity. Cell cycle analysis employs propidium iodide staining of DNA content. Apoptosis detection combines Annexin V (which binds phosphatidylserine on early apoptotic cells) with propidium iodide for membrane integrity. The DHR assay diagnoses CGD by measuring conversion of dihydrorhodamine to fluorescent rhodamine by the oxidative burst, which is absent in CGD.
Fluorescence-Activated Cell Sorting (FACS) extends flow cytometry by physically sorting cells into separate collection tubes based on their fluorescence profile, enabling isolation of pure cell populations for research or clinical use.
<image>A diagram of flow cytometry principles and data analysis. Left panel: The flow cytometer schematic shows a sample tube with cells in suspension. Cells are focused into a single stream by sheath fluid (hydrodynamic focusing). A laser beam illuminates each cell. Forward scatter (FSC) detector is positioned in line with the laser (measures cell size). Side scatter (SSC) detector is at 90 degrees (measures granularity). Multiple fluorescence detectors with bandpass filters detect different fluorophore emissions. Right panel: Data analysis examples. (A) FSC vs SSC dot plot of whole blood: three populations are gated -- lymphocytes (low FSC, low SSC), monocytes (higher FSC, moderate SSC), and granulocytes (high FSC, high SSC). (B) Gated lymphocytes displayed as CD4-FITC vs CD8-PE dot plot: four quadrants showing CD4+CD8- (Th cells), CD4-CD8+ (CTLs), CD4-CD8- (includes B cells, NK cells), and CD4+CD8+ (rare in periphery, common in thymus). Percentages are shown in each quadrant. (C) Histogram of CD4-FITC fluorescence intensity: negative population (unstained control, gray) and positive population (CD4+ cells, green peak shifted to the right).</image>
IV. Western Blot (Immunoblot)
The Western blot separates proteins by size using SDS-PAGE, transfers them to a membrane, and detects specific proteins using antibodies. The procedure involves several sequential steps. First, cells or tissue are lysed to extract proteins, which are then denatured with SDS and beta-mercaptoethanol (which reduces disulfide bonds). During SDS-PAGE, SDS coats proteins with a uniform negative charge so they migrate through the polyacrylamide gel based solely on molecular weight, with smaller proteins migrating faster. Molecular weight markers (a ladder) are run alongside the samples for reference. The separated proteins are then transferred electrophoretically from the gel to a PVDF or nitrocellulose membrane. Blocking with BSA or milk solution covers non-specific binding sites on the membrane. The primary antibody specific for the target protein is incubated with the membrane and binds the target band. A secondary antibody conjugated to an enzyme (HRP) or fluorophore binds the primary antibody. Detection using a chemiluminescent substrate (ECL) produces light emission captured on film or a digital imager, or a chromogenic substrate produces a visible band. The result is a band at the expected molecular weight that confirms the presence and size of the target protein.
The classic clinical application of Western blot was the HIV Western blot, historically used as a confirmatory test. Patient serum was tested against HIV proteins separated by gel electrophoresis, detecting antibodies against specific HIV proteins including gp160, gp120, gp41 (envelope proteins), p24 (capsid), p55, and p31 (integrase). A positive result required antibodies to at least 2 of 3 key proteins (p24, gp41, gp120/160), while an indeterminate result showed some bands but did not meet full criteria. This test has now largely been replaced by 4th generation antigen/antibody combination assays and HIV-1/HIV-2 differentiation immunoassays. In research, Western blot is used to confirm protein expression, assess protein modifications such as phosphorylation, and verify knockdown or knockout experiments.
V. Additional Immunological Techniques
Immunofluorescence microscopy uses fluorophore-conjugated antibodies to detect antigens in tissue sections or cells. In the direct format, the fluorophore-conjugated antibody binds the antigen directly. In the indirect format, a primary antibody is followed by a fluorophore-conjugated secondary antibody for signal amplification. Clinically, this technique is used for ANA pattern identification (homogeneous, speckled, nucleolar, centromere patterns on HEp-2 cells) and for direct immunofluorescence of skin biopsies in SLE, pemphigus, and IgA vasculitis.
Immunohistochemistry (IHC) uses enzyme-conjugated antibodies to detect antigens in formalin-fixed, paraffin-embedded tissue sections. It is indispensable in cancer diagnosis for identifying tumor markers such as ER, PR, HER2, Ki-67, and PD-L1, and for lymphoma classification using markers such as CD20 and CD3.
Immunoprecipitation (IP) involves an antibody binding its target protein in a cell lysate, with the antibody-antigen complex captured on protein A/G beads, washed, and eluted. It is used to study protein-protein interactions (co-immunoprecipitation) and identify binding partners, and can be combined with mass spectrometry for proteomics.
The complement fixation test is a historical assay in which patient antibodies, antigen, and complement are combined. If antibody is present, complement is "fixed" (consumed), which is detected by the failure to lyse indicator red blood cells. The Coombs test (antiglobulin test) exists in two forms. The direct Coombs test detects antibodies or complement already bound to patient red blood cells, used in diagnosing autoimmune hemolytic anemia and hemolytic disease of the newborn. The indirect Coombs test detects anti-RBC antibodies in patient serum, used for pre-transfusion crossmatching.
<image>A step-by-step diagram of the Western blot technique. Step 1 (SDS-PAGE): A gel apparatus is shown with protein samples loaded into wells. Proteins migrate through the polyacrylamide gel toward the positive electrode, separating by molecular weight (smallest at bottom, largest at top). A molecular weight ladder is shown in one lane. Step 2 (Transfer): The gel is placed next to a PVDF membrane in a transfer apparatus. An electric field transfers proteins from gel to membrane, preserving the separation pattern. Step 3 (Blocking): The membrane is incubated in blocking solution (5% milk or BSA) to cover non-specific binding sites. Step 4 (Primary antibody): The membrane is incubated with primary antibody specific for the target protein. The antibody binds only to the band containing the target. Step 5 (Secondary antibody): HRP-conjugated secondary antibody binds the primary antibody. Step 6 (Detection): ECL substrate is added; HRP catalyzes a chemiluminescent reaction. The resulting image shows dark bands at positions corresponding to the target protein. A final result image shows the membrane with a single band at the expected molecular weight (e.g., 24 kDa for p24 in HIV Western blot), compared to the molecular weight ladder.</image>
