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The normal temperature for long term antibody storage is around -20 degrees Celsius. However, when developed into rapid antibody tests, they can be kept at room temperature for prolonged periods. How is that possible?
Antibodies have no problem with room temperature. Naturally, antibodies expect to survive for many days or weeks at body temperature, which is much higher than room temperature (at least, mine is).
-20°C is recommended for very long-term storage (many months or years) of antibodies, but they are typically very stable proteins and don't denature very readily at all. Storing them at room temperature can be done for a long time -- certainly weeks, in my experience, with the only real concern being the possibility of bacterial or fungal growth if the antibodies aren't sterile.
Antibody storage guide
Please always check datasheets for specific storage recommendations. We are not able to guarantee antibodies that have not been stored correctly. With proper storage and handling, most antibodies should retain activity for months, if not years.
You can also access our most popular protocols straight from your phone with the Abcam app, which features protocols, scientific support and a suite of useful tools that are handy for any bench scientist. Learn more.
For many of our antibodies, freezing at -20°C or -80°C in small aliquots is the optimal storage condition. Aliquotting minimizes damage due to freezing and thawing, as well as contamination introduced by pipetting from a single vial multiple times. Aliquots should be frozen and thawed once, with any remainder kept at 4°C.
Upon receiving the antibody, centrifuge at 10,000 x g for 20 seconds to pull down solution that is trapped in the threads of the vial, and transfer aliquots into low-protein-binding microcentrifuge tubes. The size of the aliquots will depend on how much you typically use in an experiment. Aliquots should be no smaller than 10 μL the smaller the aliquot, the more the stock concentration is affected by evaporation and adsorption of the antibody onto the surface of the storage vial.
In most cases storage at 4°C upon receipt of the antibody is acceptable for one to two weeks. It is important to follow the recommendations on the datasheet.
Enzyme-conjugated antibodies should not be frozen at all and should instead be kept at 4°C. Freezing and thawing will reduce enzymatic activity in addition to affecting the antibody binding capacity.
Conjugated antibodies – whether conjugated to fluorochromes, enzymes, or biotin – should be stored in dark vials or wrapped in foil. Exposure to light will compromise the activity of conjugates. Fluorescent conjugates in particular are susceptible to photo-bleaching and should be protected from light during all phases of an experiment.
Antibodies of the IgG3 isotype are unique in their tendency to form aggregates upon thawing and should always be stored at 4°C.
Ascites fluid may contain proteases, and should be frozen as soon as possible after receipt.
To prevent microbial contamination, sodium azide can be added to an antibody preparation to a final concentration of 0.02% (w/v). Many of our antibodies already contain this preservative at concentrations ranging from 0.02 to 0.05%. This will be indicated on the datasheets in the storage buffer section.
When not to use sodium azide
If staining or treating live cells with antibodies, or if using antibodies for in vivo studies, be sure to use preparations that do not contain sodium azide. This antimicrobial agent is toxic to most other organisms as well: it blocks the cytochrome electron transport system.
Sodium azide will interfere with any conjugation that involves an amine group, and should be removed before proceeding with the conjugation. After conjugation, antibodies can be stored in sodium azide. The exception is HRP-conjugated antibodies which should not be stored in buffers containing sodium azide, since the same inhibits HRP. An acceptable alternative is 0.01% thimerosal (merthiolate), which does not have a primary amine.
Sodium azide can be removed from antibody solutions by dialysis or gel filtration. The molecular weight of IgG is 150,000 daltons (IgM is
600,000) the molecular weight of sodium azide is 65 daltons. A micro-dialysis unit with a cut off at 14,000 daltons will retain the antibody as the azide diffuses out.
In a beaker on a magnetic stirrer kept at 4°C, use at least a liter of cold PBS per mL of antibody and stir the dialysis unit for 6 hrs. Change the PBS twice, stirring at least 6 h for each change. If possible, all materials should be sterilized and the resulting preparation should be handled aseptically.
Repeated freeze/thaw cycles can denature an antibody, causing it to form aggregates that reduce its binding capacity.
Storing at -20°C should be adequate for most antibodies there is no appreciable advantage to storing at -80°C. The freezer must not be of the frost-free variety. These cycle between freezing and thawing (to reduce frost-build-up), which is exactly what should be avoided. For the same reason, antibody vials should be placed in an area of the freezer that has minimal temperature fluctuations, for instance towards the back rather than on a door shelf.
Some researchers add the cryoprotectant glycerol to a final concentration of 50% to prevent freeze/thaw damage glycerol will lower the freezing point to below -20°C. While this may be acceptable for many antibodies, only a small percentage of the antibodies we offer have been tested for stability in this storage condition and our guarantee only applies to antibodies stored as recommended on the datasheet. Storing solutions containing glycerol at -80°C is not advised since this is below the freezing point of glycerol. Please be aware that glycerol can be contaminated with bacteria. If adding glycerol or any cryoprotectant, care should be taken to obtain a sterile preparation.
Protein concentration and stability
Diluting antibodies to working concentration and storing at 4°C for more than a day should be avoided. Proteins in general are less susceptible to degradation when stored at higher concentrations, ideally 1 mg/mL or higher. This is the rationale for including proteins such as BSA to the antibody solution as stabilizers. The added protein also serves to minimize loss of antibody due to binding to the vessel wall. Do not add stabilizing protein to antibodies that you intend to conjugate, because they will compete with the antibody and reduce the efficiency of the conjugation.
Severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2), which has caused the COVID-19 pandemic, enters target cells through the interaction of its envelope spike protein with the primary host cell receptor angiotensin-converting enzyme-2 (ACE2), which is then cleaved by a serine protease (TMPRSS2) to allow viral fusion and entry across the cell membrane 1 . Antibodies that can bind to the spike protein have the potential to neutralize viral entry into cells and are thought to play an important role in the protective immune response to SARS-CoV-2 infection 2,3,4,5,6,7,8,9,10,11 .
To predict protection against SARS-CoV-2, it is critical to understand the quantity, quality and duration of the antibody response during different stages of COVID-19 and in the convalescent period. In this regard, assessing the level of neutralizing antibodies (NAbs) that block viral entry into cells could be a critical parameter in determining protection from SARS-CoV-2 and management of convalescent plasma therapies, which are being tested as a COVID-19 treatment option 12,13,14,15 . Defining the relationship between disease severity, other individual-specific co-morbidities and the NAb response will be critical in our understanding of COVID-19 and in tailoring effective therapies.
Currently available SARS-CoV-2 antibody tests mostly lack sufficient dynamic range and sensitivity to allow for accurate detection or determination of the magnitude of the antibody response 16 . Furthermore, potential cross-reactivity among SARS-CoV-2 specific antibodies to other endemic coronaviruses could also be confounders in these tests 17,18,19,20 , thus making them less reliable. Determining neutralization activity in patient plasma also has challenges, as these assays generally rely on live virus replication, requiring a high-level biohazard security BSL-3 level laboratory. Therefore, there is an unmet need to develop sensitive antibody and virus neutralization assays that are sufficiently robust for screening and monitoring large numbers of SARS-CoV-2 infected or convalescent subjects.
To overcome these experimental challenges, here we developed: (1) Highly sensitive bead-based fluorescent immunoassay for measuring SARS-CoV-2 specific antibody levels and isotypes, and (2) Robust SARS-CoV-2 spike protein pseudovirus to measure NAb levels in COVID-19 patient plasma. We found striking differences in total antibody levels and neutralization titers between hospitalized or severe COVID-19 patients relative to outpatient or convalescent plasma donors, which were obtained with the purpose of transfer to and treatment of patients. Significant correlations between antibody levels and neutralization titers, age and NAbs to SARS-CoV were also observed. These assays and findings have important implications for assessing the breadth and depth of the humoral immune response during SARS-CoV-2 infection and for the development of effective antibody-based therapies or vaccines.
This protocol is derived from strategies developed in our recent study characterizing the human B-cell response to influenza 1 . By this technique, it is possible for a lab experienced with the process to produce milligrams of human monoclonal antibodies (hmAbs) in as little as 28 d. This ability to express and characterize antigen-specific hmAbs is extremely useful for a variety of applications. These range from elucidating the interactions of particular antibodies and antigens to exploring basic B-cell immunology or to producing valuable therapeutics. Because of the wide epitope specificity of the antibodies produced by this method, large numbers of high-affinity antibodies can be produced quickly, yielding panels of diagnostics for rapid antigen screens.
Methods to produce hmAbs
HmAbs can be produced by several methods, including immortalization of B cells with Epstein–Barr virus 2,3 , and the production of B-cell hybridomas 4 , humanization of antibodies from other species 5 , using phage display libraries 6 or generating antibodies recombinantly from isolated single B cells 7,8 . However, the technique described herein is more suited for the rapid development of a large library of antibodies with a range of specificities against a particular immunogen. In methods requiring immortalized B-cell lines, the extensive subcloning and overall shotgun approach limit the number of useful antibodies that can be produced even over extensive periods of time 9 . Current phage display and related platforms spend extensive amounts of time identifying the few candidate antibodies present and a significant portion of these turn out to be of low affinity 9 . Although phage display technology uses fully human heavy and light chain variable genes, the heavy and light chains are randomly paired in vitro, and so are more likely to induce anaphylactic responses as foreign proteins or to be autoreactive if therapeutic uses are the goal. The mAbs generated by in vitro methods or in other species do not provide a true evaluation of the epitope specificities that humans generate in vivo, limiting the use of these techniques for applications such as epitope discovery and vaccine development or evaluation. These same applications have been hindered by technologies using immortalized B-cell lines because of the relatively few specific antibodies isolated that can be generated. Finally, for potential therapeutic applications, the Fab that is produced by phage display libraries or in other species (mice) must be cloned and fused to a human Fc backbone and expressed in a human cell line. These humanizing techniques represent a significant outlay of time and resources 9 .
Comparison to current methods to produce hmAbs
There are limitations to this method that are balanced by the advantages. The other approaches described above (Epstein–Barr virus transformation, phage display, etc.) are memory B-cell–based hmAb technologies that allow a retrospective evaluation of the entire history of previous antigen exposures. This allows mAbs to be isolated even 80 years after exposure to the pathogen as recently illustrated by the cloning of antibodies against the 1918 influenza pandemic strain from a 95-year-old donor 10 . These methods negate the need for obtaining fresh samples (frozen peripheral blood mononuclear cells (PBMCs) are suitable for use) and avoid the logistical difficulties of obtaining B cells from people with active immune responses. Conversely, the power of the ASC-based hmAb approach derives from that very limitation: the approach relies on isolating activated plasmablasts at the peak of the immune response such that the majority of the hmAbs isolated are antigen specific. Thus, although a human vaccine must be available, as well as a donor to receive the vaccination and donate blood, this allows an unprecedented efficiency to generate many specific mAbs. In addition, the process provides a window directly into ongoing immune responses. For example, we have observed expansions of the ASC population during natural infections (unpublished observations). Therefore, it is likely that the procedure can be used to make antibodies from ASCs induced during or soon after natural infections, or to make anti-self antibodies from patients with certain autoimmune disorders. Finally, our method as described herein relies on transient transfection for production of the antibodies, allowing rapid screening of many antibodies but making large-scale antibody preparations difficult. However, should production need to be scaled up, methods of producing stable transfectants and manipulation of the production cell lines could easily be adapted.
The steps intrinsic to our process of producing recombinant hmAbs are a modification of a system that has been used to elucidate basic mechanisms of B-cell immunology and autoimmunity by both us 11,12 and others 7,8 . However, the protocol described here is the first integration of this process that yields high-affinity hmAbs specific to an antigen of interest relatively quickly and critically, independent of any specific antigen staining. Thus far, this technique has produced anti-influenza, anti-anthrax toxin antibodies and anti-pneumoccocal hmAbs and can be adapted easily to any immunogen with which humans are vaccinated. For example, another group has generated anti-tetanus antibodies using variable genes from ASCs that were expressed in Escherichia coli 13 . This versatility can provide a library of human antibodies against targets that require a rapid response, such as bioterrorism agents or viral epidemics, or large libraries of hmAbs such as in vaccine evaluation.
Potential applications of the hmAbs produced by vaccinating antigens
There is a common misconception that active immunization precludes the need for passive immunization. Even though a vaccine is necessary to produce antibodies by the method we describe, the monoclonal antibodies produced by our system could be used as passive immunotherapy treatments in a large variety of cases. Pooled human immunoglobulin is currently used as a treatment for several agents, including hepatitis B, tetanus and rabies 14 . Pooled sera carry risks, including potential anaphylactic responses or autoimmune reactions, that could be avoided by a single effective neutralizing monoclonal antibody. Furthermore, in cases of a bioterrorist attack with a pathogen such as anthrax that the general public is not vaccinated against, hmAbs could provide rapid protection against the pathogen while the antibiotics begin to decrease the bacterial load. Similarly, monoclonals against toxins such as botulinum neurotoxin could aid in treating those exposed, as active immunization with vaccine will take at least 2 weeks to confer protection. Finally, there are many immunocompromised populations in which vaccines are ineffective 15 . In these cases, including the elderly and the very young, monoclonal passive immunotherapies could be crucial in treating infectious diseases. A final potential application is for the development of therapeutic antibodies to treat chronic or antibiotic-resistant infectious diseases. Some key examples include the substantial effort now being invested to isolate the rare broadly neutralizing antibodies that can control various strains of HIV 16 . Although these reagents could be used directly as an adjunct to antiviral drugs in controlling viremia, the more important application may be the ability to evaluate many of these antibodies to understand how a vaccine can elicit them. A second example is the potential to generate therapies against antibiotic-resistant bacteria directly from patients that are clearing the infection: new antibiotics are rare, but antibodies can clear these infections despite drug resistance 17 . Neutralizing antibodies from these patients could be used directly or pharmaceutical targeting of the neutralizing epitopes discovered could substantially increase our treatment options. In all of these cases, primarily by the ability to isolate many specific hmAbs rapidly, our technique greatly increases the potential for using monoclonal antibody therapeutics for a wide variety of infectious diseases and bioterrorist agents.
A flow chart briefly describing all stages in this protocol is shown in Figure 1.
Under optimal conditions, an experienced laboratory can complete the entire procedure from vaccination to antibody in as little as 28 d.
In this protocol, antibody-secreting cells (ASCs) are first isolated from whole blood collected 7 d after vaccination with an immunogen. We have successfully made antibodies following vaccination with Fluvirin (2005–2006, 2006–2007 and 2007–2008), Pneumovax23 and Biothrax. PBMCs are isolated using a standard lymphocyte separation protocol. The frequency of antigen-specific ASCs is analyzed using a standard ELISpot protocol 18 (see Box 1). This assay enumerates the number of IgG-producing ASCs, as well as antigen-specific ASCs. The percentage of antigen-specific, IgG-producing ASCs is a useful measure of the donor's response to the vaccine and therefore the approximate quantity of high-affinity antibodies produced.
The cells are then sorted by flow cytometry. First, the live cell gate, including larger blasting cells, is set using forward versus side scatter. The ASCs are bulk sorted by first gating on CD19 high /CD20 low to neg /CD3 neg and then on CD27 high /CD38 high cells as shown in Figure 2. The appropriate IgG, IgM and IgD gates are set to obtain IgG-producing ASCs, although it is also possible to use this method to isolate IgM-producing ASCs as well. Finally, the purified ASCs are single cell sorted into single cell PCR plates loaded with catch buffer containing RNase inhibitor.
First, the live cell gate is set, including blasting cells, then CD19 high /CD20 low to neg /CD3 neg and CD27 high /CD38 high . Finally, appropriate IgG IgM, and IgD gates are set to obtain the precise population of interest, improving the immunoglobulin constant region-specific priming efficiency.
Using both RT-PCR and nested PCR, the antibody genes in each cell are amplified on a per cell basis. The RT-PCR is accomplished using a cocktail of nine primers, designed to cover all of the families of variable (V) genes possible (Table 1). The nested PCR is performed to amplify the DNA enough to obtain sequences of the heavy and light chain V genes. This is necessary for the cloning PCR. In this step, highly specific primers for each V gene family are used to amplify the DNA for cloning. The 'cloning PCR' primers are designed both to incorporate the cloning restriction sites and to place the VDJ heavy or VJ light chain genes in frame with the signal peptide sequences and constant region genes within the respective cloning vectors. Cloning sites were incorporated into the vectors that are specific for the particular heavy or light chain vectors to allow proper, in-frame incorporation of the variable gene inserts. The inserts and vectors are then digested and purified for cloning. The heavy and light chain DNA from each single cell is then cloned into separate vectors and transformed. At least four colonies from the transformation are grown, mini-prepped and sequenced. The sequences from each colony are compared and the colony most closely matching the consensus is then chosen for further amplification to maxi scale.
Transiently transfected human kidney epithelial cells (the HEK293 cell line 19 ) are used to produce the antibody. Polyethyleneimine-based transfection is used with equimolar amounts of heavy and light chain vector according to standard protocols 20 . The cells are allowed to produce antibody for 5 d. The transfection media containing the hmAbs are then purified using protein A agarose beads and concentrated using commercial protein concentrators. During the final stage, the hmAbs are analyzed for concentration, purity and reactivity.
Cell Lysis and Immunoprecipitation with CHAPS Detergent
The CHAPS immunoprecipitation buffer (CIB-1) is formulated to maintain intermolecular interactions following transmembrane and cytosolic protein extraction. The CHAPS buffer is ideal for pull down co-immunoprecipitation (i.e.Co-IP) analysis, since it maintains protein conformation as well as protein complex assembly.
Following cell lysis and cellular protein solubilization into CHAPS buffer, assembled proteins can be isolated together using immunoprecipitation technique. An immunoprecipitating antibody is added to bind to one of the assembled protein targets. A solid support, such a Protein-G Sepharose, is then added to non-covalently link and bind to the antibody-protein complex and make it insoluble. Following low-speed centrifugation, sedimentation and wash, the solid support-antibody-protein complex is isolated together.
Proteins assembled with the protein targeted by the immunoprecipitating antibody become co-immunoprecipitated. By developing Western blots with a different antibody targeting a co-assembled protein, the researcher can characterize and quantify the interaction among two or more protein species. Functional biochemical activity assays can also be employed to characterize co-immunoprecipitated and assembled biomolecules.
- Microcentrifuge at 4°C
- Rocker table
- Cold room or refrigerated case
- Antibody suitable for immunoprecipitation as specified by the vendor. Generally, the immunoprecipitating antibody must recognize the NATIVE conformation.
- Western blots applications: Antibody suitable for Western blot detection as specified by the vendor. This antibody must recognize the DENATURED state of the targeted protein.
Suggested Controls and Samples for Comparisons
- Perform immunoprecipitation protocol with an irrelevant antibody to control for appearance of spurious protein bands.
- Resolve in Western blots the original unfractionated cell lysate, the unbound fraction (e.g. the fraction containing proteins not bound to the protein-A or G solid support) and the immunoprecipitated materials to confirm fractionation and enrichment.
- Develop a Western blot that detects the protein directly targeted by the immunoprecipitating antibody. This procedure confirms that immunoprecipitation occurred.
A. Protocol for Cell Culture
It is recommended that cells are cultured to 80-90% confluency prior to performing cell lysis and immunoprecipitation. Cells should be washed free of serum proteins using PBP prior to performing immunoprecipitation to prevent appearance of non-specific serum protein bands in downstream Western blots.
B. Protocol for Cell Lysis with CHAPS detergent
We recommend using 300 uL of CHAPS solution for one to three 10 cm cell culture dishes of lysed cells. Scale accordingly for other numbers or sizes of cell culture dishes according to the surface area of the dish. Prior to lysis, make the CHAPS Buffer ice cold and add protease and phosphatase inhibitors.
Cells should be washed free of serum proteins using PBS prior to performing immunoprecipitation to prevent appearance of non-specific serum protein bands in downstream Western blots. Remove the cell culture medium and gently wash the 10cm cell culture dish with 5 mL PBS, three times. Use a gentle stream of PBS to prevent excessive loss of cells from the plate. After the washes, cover the cell culture dish with the lid and place the cell culture dish on a bed of ice
- Lyse cells and generate a supernatant fraction rapidly as follows: Dispense 300 uL ice cold CHAPS with protease/phosphatase inhibitors over the cell layer, rotate the plate by hand to cover cells with a film of CHAPS, then immediately dislodge the cells with a cell scraper. Use a transfer pipette with a wide opening to siphon the cells into a 1.5 mL microcentrifuge tube. The 300 uL Chaps solution can also be transferred sequentially to up to three separate 10 cm plates of cells before the cell suspension is placed into a 1.5 mL microcentrifuge tube.
- Place the 1.5 mL microcentrifuge tube with cell suspension on ice for 10 min: Strongly tap the tube several times during this 10 min period to facilitate cell membrane dissolution. You can also use a rocker table to rotate the cell suspension to further facilitate cell membrane dissolution. Do not vortex the cell lysate if immunoprecipitation is planned.
- Centrifuge the cell lysate in a cooled microcentrifuge at full speed for 15 min to partition supernatant and pellet. Collect the supernatant fraction, which contains extracted membrane and cytosolic proteins, and dispense this supernatant into another 1.5 mL microcentrifuge tube that is placed in ice. This transferred supernatant corresponds to the unfractionated supernatant fraction. Set aside a small aliquot for comparisons to the immunoprecipitated materials that are generated later in the protocol. The unfractionated supernatant can be stored at -20°C, or -80°C for longer term storage.
C. Protocol for Immunoprecipitation with CHAPS detergent
- Add immunoprecipitating antibody to the unfractionated supernatant fraction, using the antibody titer recommended by the manufacturer. Place the tube with the immunoprecipitation reaction on a rocker table under refrigeration (such as a cold room, or refrigerated case) for 15-30 min to mix the antibody-supernatant mixture. You may opt to initially try the shorter 15 min period for immunoprecipitation, a time period that is more apt to maintain low affinity interactions.
- Directly dispense 60 uL of immunoprecipitation beads (e.g. Sepharose-G beads, or protein-A beads) into every 300 uL to 2 mL aliquot of unfractionated supernatant with immunoprecipitating antibody. Place this tube on a rocker table for another 30 min to 1 hr under refrigeration to generate an insoluble solid support–antibody-protein complex.
- Use a refrigerated microcentrifuge at 3000 rpm for 5 minutes to sediment the immunoprecipitation beads. The resulting supernatant is the unbound fraction. Collect the unbound fraction without disturbing the immunoprecipitating bead pellet. Store the unbound fraction at -20°C or -80°C.
- Add an additional aliquot of fresh 300 uL ice cold CHAPS with protease and phosphatase inhibitors to the immunoprecipitation beads. Gently rotate the tube 180° by hand three times and centrifuge again at 3000 rpm for 5 minutes. Remove and discard the wash supernatant. Repeat this wash procedure two more times. After the last wash, use a microcentrifuge to sediment the immunoprecipitation beads at 14000 rpm for 15 min. Remove as much of the wash solution as possible using a pointed plastic Pasteur pipette. The immunoprecipitated fraction, which is bound to the beads, becomes sedimented at the bottom of the tube.
D. Protocol for Elution of the Immunopreciptated Fraction from the Beads and Western Blot with CHAPS detergent
- There are several methods to elute the immunoprecipitated proteins from the solid support to release a soluble immunoprecipitated fraction. The simplest method applicable for subsequent Western blotting is to apply Laemmli Sample Buffer (LSB with mercaptoethanol) directly to the immunopreciptation beads: Add 100l LSB to each 60 uL of sedimented immunoprecipitation beads. Vortex the LSB – immunoprecipitate solution at full speed for 30 sec, and then heat at 60°C for 10 min. Now sediment the beads using low speed centrifugation (3000 rpm) for 5 min. Collect the supernatant The immunprecipitated proteins (along with the immunoprecipitating antibody) will be released into the supernatant.
- The supernatant can be resolved in subsequent Western blots. Alternatively, if your protein of interest aggregates easily when heated at high temperature in LSB, heat instead at 37°C for 30 min. and follow the same aforementioned steps. Resolve in consecutive gel lanes and Western blots, 1) the unfractionated fraction, 2) unbound fraction and 3) immunoprecipitated fraction. With successful immunoprecipitation, you should observe enrichment of your protein of interest, along with any co-assembled proteins in the immunopreciptated fraction.
E. Protocol for Tissue Homogenization Prior to Immunopreciptiation with CHAPS detergent
This process should be performed on ice.
- Pulverize approximately 90 uL of tissue. Place tissue in a 1.5 mL round bottom microcentrifuge tube.
- Add general phosphatase and protease inhibitor cocktails to 500 uL of ice-cold CHAPS Lysis and Immunoprecipitation Buffer
- Add 500 uL CHAPS buffer with inhibitors to pulverized tissue.
- Homogenize tissue with a mini pestle-homogenizer using 15 strokes, 3 seconds/stroke on ice.
- Centrifuge 12,000 g for 15 min at 4°C
- Remove supernatant (without lipid layer) and transfer into another 1.5 mL tube
- Centrifuge again at 12,000 g for 15 min at 4°C
- Transfer supernatant to another tube. The supernatant fraction contains the extracted proteins that can be used for immunoprecipitation.
- Immunoprecipitation did not occur. Resolution: 1) You may have to empirically identify an antibody for immunoprecipitation that recognizes the native conformation of the epitope. 2) The protein complex is not maintained outside of a live-intact cell. Cross-linking procedures using membrane permeable cross-linking reagents and live cells may be required to capture the interaction and assembly among proteins.
- The IgG heavy chain of the immunoprecipitating antibody (that was dispensed in the immunoprecipitation solution) migrates in gels at the same position as a protein of interest, and therefore masks its appearance in a Western blot. Resolution: For Western blot development, use an antibody derived from an animal host different from the immunoprecipitating antibody. In this case, the appropriate secondary-HRP conjugated antibody will not bind appreciably to the immunoprecipitating antibody that was transferred to the Western blot, preventing the appearance of an overlaying Western blot band.
Animal efficacy data shows robust immune response and safety
Researchers tested two vaccine candidates of AAVCOVID that were qualitatively different in how they would elicit an immune response. They tested each candidate in mouse and nonhuman primate models at different doses to determine safety and immunogenicity (the ability to produce an immune response).
The immunogenicity following a single intramuscular injection was shown to be potent in inducing neutralizing antibodies in two mouse strains and both genders, a mouse model for aging and an obese mouse model. Both vaccine candidates demonstrated durable responses from a single dose administration for at least three months. Early onset of antibody responses were seen by day 14 and rates increased over time.
In non-human primates, AAVCOVID candidates were shown to retain peak immunogenicity for at least five months following a single-dose injection. Responses in both animal models were complemented by functional memory T-cell responses, important to confer durable immunity.
Mice and nonhuman primates tolerated the vaccine candidates well with no adverse safety events observed. While these results need to be confirmed in human studies in healthy volunteers, the researchers hope the favorable safety profile of established AAV-based gene therapies already approved for use in Europe and the U.S. -- which are often given at much higher doses in patients -- will translate to the vaccine.
Analysis of Results: Fluorescent vs. Light Microscopy
Both immunofluorescent ICC and IHC require an understanding of the configuration of epifluorescence or confocal microscope that will be used to analyze the sample. The best results will be obtained when spectral characteristics of the fluorophores conjugated to the primary or secondary antibody are matched to the excitation source (usually a laser) and emission filters of the available microscope. For more about designing a fluorescence microscopy analysis experiment, visit this page.
White light or brightfield microscopy may be more accessible to researchers as the necessary equipment is available in most labs, but is limited by the number of targets which may be simultaneously detected in the same tissue section. This is because deposition of the chromogen at the site of antibody binding to antigen in the tissue is dependent on enzyme-substrate activity, and the most common, reliable reagents (such as diaminobenzidine, or DAB, and alkaline phosphatase, or AP) do not offer multispectral colorimetric detection that would permit simultaneous detection of multiple targets.
We thank Prof. H. Eric Xu (Shanghai Institute of Materia Medica) for providing RdRp protein. We also thank Healthcode Co., Ltd., Hangzhou Bioeast biotech Co., Ltd. and Vacure Biotechnology Co.,Ltd. for providing the proteins. This work was supported by grants from the National Mega-Projects of Science Research for the 13th Five-year Plan of China (No. 2018ZX10302302002-001), the Natural Science Foundation of China (No. 81971909), and the Fundamental Research Funds for the Central Universities (HUST COVID-19 Rapid Response Call No. 2020kfyXGYJ040) to X-L Fan. This work was also partially supported by National Key Research and Development Program of China Grant (No. 2016YFA0500600), Interdisciplinary Program of Shanghai Jiao Tong University (No. YG2020YQ10), National Natural Science Foundation of China (No. 31900112, 21907065, 31970130 and 31670831) to S-C Tao.
Product DetailsPreservative Stabilisers
|0.09%||Sodium Azide (NaN3)|
|1%||Bovine Serum Albumin|
|0.09%||Sodium Azide (NaN3)|
|1%||Bovine Serum Albumin|
|Annexin V:APC||200 tests|
|Propidium Iodide Staining Solution||2X 100 tests||10X Binding Buffer||100 ml|
|Annexin V:PE||200 tests|
|7-AAD Viability Staining Solution||2X 100 tests||10X Binding Buffer||100 ml|
Interim Guidance for Antigen Testing for SARS-CoV-2
Antigen tests are commonly used in the diagnosis of respiratory pathogens, including influenza viruses and respiratory syncytial virus. The U.S. Food and Drug Administration (FDA) has granted emergency use authorization (EUA) for antigen tests that can identify SARS-CoV-2. See FDA&rsquos list of In Vitro Diagnostics EUAs external icon .
Antigen tests are immunoassays that detect the presence of a specific viral antigen, which implies current viral infection. Antigen tests are currently authorized to be performed on nasopharyngeal or nasal swab specimens placed directly into the assay&rsquos extraction buffer or reagent. The currently authorized antigen tests include point-of-care, laboratory-based, and self-tests, and they are applicable to people of any age. See Table 1 for additional information about antigen tests.
Antigen tests are relatively inexpensive, and most can be used at the point of care. Most of the currently authorized tests return results in approximately 15&ndash30 minutes. Antigen tests for SARS-CoV-2 are generally less sensitive than real-time reverse transcription polymerase chain reaction (RT-PCR) and other nucleic acid amplification tests (NAATs) for detecting the presence of viral nucleic acid. However, NAATs can remain positive for weeks to months after initial infection and can detect levels of viral nucleic acid even when virus cannot be cultured, suggesting that the presence of viral nucleic acid may not always indicate contagiousness.
Proper interpretation of both antigen test results and NAATs (when indicated) is important for accurate clinical management of patients or people with suspected COVID-19, or for identification of infected people when used for screening.
The clinical performance of diagnostic tests largely depends on the circumstances in which they are used. Both antigen tests and NAATs perform best if the person is tested when their viral load is generally highest. Because antigen tests perform best in symptomatic people and within a certain number of days since symptom onset, antigen tests are used frequently on people who are symptomatic. Antigen tests also may be informative in diagnostic testing situations in which the person has a known exposure to a person with COVID-19.
Accumulation of data on the performance of antigen tests in different situations has helped guide the use of these tests as screening tests in asymptomatic people to detect or exclude SARS-CoV-2 infection. See FDA&rsquos Recommendations for healthcare providers using SARS-CoV-2 diagnostic tests for screening asymptomatic individuals for COVID-19 external icon . Also see information from the Centers for Medicare & Medicaid Services (CMS) on Updated CLIA SARS-CoV-2 Molecular and Antigen Point of Care Test Enforcement Discretion external icon .
Antigen tests have been used for screening testing in high-risk congregate housing settings, such as nursing homes, in which repeat testing has quickly identified people with COVID-19, informing infection prevention and control measures, thus preventing transmission. In this case, and where rapid test turnaround time is critical, there is value in providing immediate results with antigen tests, even though they may have lower sensitivity than NAATs.
Healthcare providers and public health practitioners should understand test performance characteristics to recognize potentially false negative or false positive test results and to guide additional confirmatory testing and management of the patient or person. Laboratory and testing professionals who perform antigen tests should understand the factors that affect the accuracy of antigen testing, as described in this guidance. Healthcare providers, laboratory and testing professionals, and public health practitioners should also understand the differences among diagnostic, screening, and surveillance testing. See CDC&rsquos Overview of Testing for SARS-CoV-2, and Testing Strategies for SARS-CoV-2. Also see FDA&rsquos FAQs on Testing for SARS-CoV-2 external icon .
Regulatory Requirements for Using Antigen Tests for SARS-CoV-2
FDA regulates in vitro diagnostic devices and has provided recommendations and information regarding EUA requests for COVID-19 diagnostic tests in the Policy for Coronavirus Disease-2019 Tests During the Public Health Emergency (Revised) (&ldquoPolicy for COVID-19 Tests&rdquo) external icon and the EUA templates referenced in that policy. COVID-19 tests and test systems used for diagnostic or screening testing, including those for antigen testing, must have received an EUA from FDA or be offered under the policies in FDA&rsquos Policy for COVID-19 Tests external icon . Every antigen test for SARS-CoV-2 authorized for use by FDA is included on FDA&rsquos list of In Vitro Diagnostics EUAs external icon . The intended use of each test, available in the Instructions for Use and in the Letter of Authorization, defines the population in which the test is intended to be used, the acceptable specimen types, and how the results should be used.
Laboratory and testing professionals who conduct diagnostic or screening testing for SARS-CoV-2 with antigen tests must also comply with Clinical Laboratory Improvement Amendments (CLIA) regulations. Any laboratory or testing site that intends to report patient-specific test results to a person or healthcare provider must first obtain a CLIA certificate and meet all requirements to perform that testing. For more information, see CMS&rsquo How to Obtain a CLIA Certificate external icon . CMS has provided additional information on Enforcement discretion for the use of SARS-CoV-2 point-of-care testing on asymptomatic individuals external icon .
Performance of Antigen Tests for SARS-CoV-2
It is important for healthcare providers and testing personnel to understand the performance characteristics, including sensitivity, specificity, and positive and negative predictive values, of the particular antigen test being used, and to follow the manufacturer&rsquos instructions for use, which summarize performance characteristics. See FDA&rsquos In Vitro Diagnostics EUAs external icon for detailed information about specific authorized tests.
The &ldquogold standard&rdquo for clinical diagnostic detection of SARS-CoV-2 remains laboratory-based (moderate- and high-complexity) NAATs. Thus, it may be necessary to confirm an antigen test result with a laboratory-based NAAT, especially if the result of the antigen test is inconsistent with the clinical context. Table 1 summarizes the differences between NAATs and antigen tests. Clinical performance of NAATs and antigen tests may differ from clinical utility when considering issues of test availability, quality of specimen collection and transport, and turnaround times of results. Based on their instructions for use, some point-of-care NAATs may not be used for confirmatory testing. NAATs that generate presumptive results are not appropriate for use in confirmatory testing.
The sensitivity of antigen tests varies but is generally lower than most laboratory-based NAATs. The antigen level in specimens collected either before symptom onset, or late in the course of infection, may be below the tests&rsquo limit of detection. This may result in a negative antigen test result, while a more sensitive test, such as most NAATs, may return a positive result. Studies external icon have shown that antigen tests have comparable sensitivity to laboratory-based NAATs when viral load in the specimen is high and the person is likely to be most contagious.
The specificity of antigen tests is generally as high as most NAATs, which means that false positive test results are unlikely when an antigen test is used according to the manufacturer&rsquos instructions. Despite the high specificity of antigen tests, false positive results will occur, especially when used in communities where the prevalence of infection is low &ndash a circumstance that is true for all in vitro diagnostic tests. In general, for all diagnostic tests, the lower the prevalence of infection in the community, the higher the proportion of false positive test results.
Positive and negative predictive values of all in vitro diagnostic tests (e.g., NAAT and antigen tests) vary depending upon the pretest probability. Pretest probability considers both the prevalence of the target infection in the population that is being tested as well as the clinical context of the individual being tested. If the prevalence of infection in the community is high, and the person being tested is symptomatic, then the pretest probability is generally considered high. If the prevalence of infection in the community is low, and the person being tested is asymptomatic and has not had any known contact to a person with COVID-19, then the pretest probability is generally considered low. See CDC&rsquos Interpreting Results of Diagnostic Tests for additional information on the relationship between pretest probability and the likelihood of positive and negative predictive values.
To help estimate pretest probability, CDC recommends that laboratory and testing professionals who perform antigen testing determine infection prevalence based on a rolling average of the positivity rate of their own SARS-CoV-2 testing over the previous 7&ndash10 days. Alternatively, state health departments generally publish COVID-19 data on testing positivity rates and case rates for their communities.
Processing of Antigen Tests for SARS-CoV-2
The Conditions of Authorization in the antigen EUAs specify that CLIA-certified laboratories and testing sites are to follow the manufacturer&rsquos instructions for use, typically found in the package insert, when performing the test and reading test results. The authorized instructions for use for each test can also be found at FDA&rsquos In Vitro Diagnostics EUAs external icon .
For example, the performance of antigen tests can be affected if the test components are not stored and handled properly. They should never be frozen and should always be allowed to reach room temperature (15-30°C) before use. The package insert for these tests includes instructions for handling of the test cartridge/card, such as ensuring it remains in its sealed pouch until immediately before use.
The package insert for antigen tests also includes instructions about how to read the test results, including the appropriate time to read the results and whether the results should be interpreted visually or with an instrument analyzer. Reading the test before or after the specified time could result in false positive or false negative test results.
Processing multiple specimens successively or in batch mode may increase the risk of contamination and may make it more challenging to ensure that each specimen is incubated for the correct amount of time before the result is read. Refer to the package insert for the correct incubation time for that test, and then monitor and ensure proper timing for each specimen during testing and when reading results.
All testing for SARS-CoV-2, including antigen testing, depends on the integrity of the specimen, which is affected by procedures for both specimen collection and handling. Improper specimen collection, such as swabbing the nostril too quickly, may cause insufficient specimen collection, resulting in limited amounts of viral genetic or antigenic material for detection. Time from specimen collection to testing should be minimized, and the temperature of the specimen during this time must be controlled. See CDC&rsquos Interim Guidelines for Collecting, Handling, and Testing Clinical Specimens for COVID-19.
Quality assurance procedures should be followed to prevent cross-contamination and inaccurate test results. For example, users should follow the manufacturer&rsquos instructions, as well as state and local guidance, for when and how often to perform testing on control specimens. If antigen testing returns multiple unexpected positive results, it may be appropriate to stop testing patient specimens, review all procedures, disinfect all surfaces, change gloves, and run control specimens before restarting the testing of patient specimens. In such circumstances, confirmatory testing should be considered for people who received unexpected results, regardless of pretest probabilities.
Decontaminate work surfaces and equipment with appropriate disinfectants by using an EPA-approved disinfectant for SARS-CoV-2, following the manufacturer&rsquos recommendations for use, such as dilution, contact time, and safe handling. See EPA&rsquos List of Disinfectants for COVID-19 external icon . Gloves should be changed before collecting, handling, and processing a new specimen in the antigen test system. Failing to change gloves can increase the risk of cross-contamination and false antigen test results. See CDC&rsquos guidance on Point-of-Care Testing, and Interim Laboratory Biosafety Guidelines for Handling and Processing Specimens Associated with Coronavirus Disease 2019 (COVID-19).
Some antigen tests have explored the use of viral transport medium (VTM) during specimen collection, but the use of VTM may cause false test results from either cross-reactivity with the capture antibodies or dilution of the specimen that decreases the sensitivity of the test. Laboratories and testing sites should refer to the instructions for use and the package insert that are specific for the test that they are using regarding the use of VTM.
Also see FDA&rsquos Letter to Clinical Laboratory Staff and Health Care Providers external icon on the potential for false positive results with antigen tests, and CDC&rsquos guidance on Point-of-Care Testing.
Evaluating the Results of Antigen Testing for SARS-CoV-2
Evaluating the results of an antigen test for SARS-CoV-2 depends primarily on the clinical and epidemiological context of the person who has been tested (e.g., symptoms, exposure to others with COVID-19, vaccination status, previous infection status, or setting in which they live). For additional details on testing recommendations see guidance for fully vaccinated people. A particularly important aspect of epidemiological context is whether the person to be tested is a resident or an employee of a congregate living facility. In addition, evaluating the results of an antigen test for SARS-CoV-2 should consider the performance characteristics (e.g., sensitivity, specificity) and the instructions for use of the FDA-authorized test, and the prevalence of SARS-CoV-2 infection in that particular community (percent positivity rate over the previous 7&ndash10 days or the number of cases in the community relative to the population size).
The evaluation of an antigen test result should consider whether the person has experienced symptoms, and if so for how long. Generally, healthcare providers can rely upon a positive antigen test result for a symptomatic patient because the specificity of current FDA-authorized antigen tests is high.
The sensitivity of current FDA-authorized antigen tests varies, and thus negative diagnostic testing results should be handled depending on the circumstances. In most circumstances, the manufacturers&rsquo instructions for use of antigen tests indicate that negative test results should be considered &ldquopresumptive,&rdquo meaning that they are preliminary results. See FDA&rsquos In Vitro Diagnostics EUAs external icon .
It may be appropriate to confirm antigen test results with a laboratory-based NAAT. For confirmatory testing, CDC recommends using a laboratory-based NAAT that has been evaluated against the FDA reference panel for analytical sensitivity. See FDA&rsquos SARS-CoV-2 Reference Panel Comparative Data external icon . NAATs that generate presumptive results are not appropriate for use in confirmatory testing.
CDC has developed two general antigen testing algorithms to accommodate two broad categories of use for antigen tests. CDC recommends following one of these two antigen testing algorithms to determine when confirmatory testing is recommended.
The first algorithm is designed for those who live in congregate settings, such as long-term care facilities, correctional and detention facilities, homeless shelters, and other group shelters. In these settings, correct case identification is particularly important because of the need to group isolated people together or in close proximity, so false positive test results can have significant negative consequences. See Figure 1, also available as a PDF. This algorithm is not designed for employees of congregate living facilities, who can quarantine and isolate outside of the facility if necessary.
The second algorithm is designed for community testing among people who do not live in congregate settings. The primary objective of this testing is to reduce the transmission of SARS-CoV-2 in the community, where there are concerns for introduction and widespread transmission, by quickly identifying and isolating people who are infected. See Figure 2, also available as a PDF.
How To Optimize Your ELISA Experiments
The enzyme-linked immunosorbent assay (ELISA) is one of the most sensitive and reproducible technologies available. These assays are rapid, simple to perform, and easily automated. As with any assay, the reproducibility and reliability of ELISAs depend upon proper technique and attention to detail.
The ELISA technique is divided into:
Direct ELISA, where the antigen is immobilized on the ELISA plate, and the primary antibody carries the label (Figure 1 A)
Indirect ELISA, where the antigen is immobilized on the ELISA plate, and the secondary antibody carries the label (Figure 1 B)
Sandwich ELISA, where two primary antibodies (for capture and detection) embed the antigen, forming a "sandwich" and then the complex is recognized by a secondary labelled antibody (Figure 1 C).
Figure 1. Types of ELISA formats: A) direct, B) indirect and C) Sandwich ELISA.
Matched antibody pair in ELISA kit
Matched pair refers to sets of antibodies that are known to recognize different epitopes on the same protein antigen, so they can be used together for the capture and detection of a single antigen in a sandwich ELISA. The antibodies used in ELISA assays can be monoclonal, polyclonal, or a combination of both.
Monoclonal antibodies can be used for all antibody-containing steps in all types of ELISAs. They are commonly used in sets as matched pairs in sandwich ELISAs but can be used for capture or detection in conjunction with a polyclonal antibody to enhance signal or to provide a greater chance of capturing an antigen from a complex solution.
Due to the heterogeneity of antibodies present in polyclonal antibody solutions and the wide representation of epitopes present, polyclonal antibodies can be powerful tools for the detection of an antigen, often yielding higher signal levels than monoclonal antibodies. However, polyclonal antibodies are more likely to share one or more epitopes with closely related proteins, resulting in higher non-specific signal. One method of reducing this problem is to use affinity purified or cross-absorbed polyclonal antibodies. To increase assay sensitivity, the detection method for an ELISA can be switched from direct to indirect detection using a polyclonal antibody.
A variety of samples can be tested in an ELISA, and the choice of assay conditions will depend upon the complexity of the sample and the expected amount of antigen present.
It is important to test all samples in duplicate or triplicate in conjunction with a known standard to ensure the accuracy of results and for quantitation.
It is better to test several dilutions of a sample to make sure the final results fall within the linear portion of the standard curve. Highly concentrated samples can underestimate concentration while highly diluted samples can overestimate concentrations.
ELISA blocking and washing steps
Blocking is often necessary to prevent non-specific binding of detection antibodies to the multi-well plate surface itself. When a plate is fully blocked, assay sensitivity will be enhanced since non-specific signal will be reduced.
A thorough washing procedure is essential for obtaining reliable ELISA results. It is important to completely aspirate liquid from all wells by gently lowering an aspiration tip into the bottom. Avoid scratching the inside of the well. When washing is complete, it is recommended to invert the plate and dry it on absorbent tissue.
How to optimize your ELISA experiment
Although each component is described separately, in many cases it is possible to optimize two components simultaneously by performing a checkerboard titration.
1) Capture Antibody Concentration
2) The Blocking Buffer
- Prepare different blocking solutions. If the blocking solution is not preformulated (i.e., it is a single protein, such as BSA), try different concentrations of the protein.
3) The Standard Diluent
Try to match the standard diluent as closely as possible to the matrix of the sample.
If the matrix itself cannot be exactly duplicated then test different standard diluent solutions and check the standard curve and linearity of dilution for the sample. It may be necessary to choose a different diluent.
4) Sample Concentration
- Prepare different concentrations of the sample, keeping in mind the detection limit of the substrate. To confirm that the biological sample matrix is not masking or enhancing the signal, spike-and-recovery and linearity-of dilution experiments should be performed.
5) The Detection Antibody Concentration
6) The Enzyme Conjugate Concentration
- Prepare different concentrations of the enzyme conjugate according to the ELISA kit range described for the substrate.
7) Signal Detection
Select substrate(s) based on the amount of the antigen in the sample and ability to detect it with a plate reader.
If the antigen can clearly be detected then the substrate is appropriate.
If the antigen is below the threshold for detection then select a more sensitive substrate.
Common ELISA issues
1) Weak or no signal in ELISA
It is recommended that all reagents are at room temperature for 15–20 minutes before starting the assay.
Incorrect storage of components. Double-check storage conditions on the kit label. Most kits need to be stored at 2–8 o C.
Reagents added/prepared incorrectly. Check the protocol, ensure reagents were added in the proper order, and prepared to correct dilution.
Overexposure of fluorescent reagents. Intense light can cause photo-bleaching by decomposition of the fluorophore. Protection of the fluorophore from light is essential for effective signal generation at the end of the assay.
Components require further optimization. One of the components of the assay may be at a limiting concentration, resulting in lower overall signal.
Degradation of reagents. One of the reagents may have degraded or been contaminated, in which case it should be replaced.
Antibodies are an inefficient pair or lack sufficient affinity towards the target. The antibodies used may not bind effectively to the antigen or may not work in combination with each other. In such cases, alternative antibodies must be tested.
Detection system requires further optimization. The detection system (substrate) may not be sensitive enough to give the signal, or the standard curve may not be appropriate for the sample. It might be necessary to concentrate the sample or switch to a more sensitive substrate.
2) High background
Insufficient washing or blocking. High background signal is commonly the result of either insufficient washing or blocking, sample components or antibodies cross-reacting with the blocking buffer, or the use of too much enzyme conjugate.
It is important to balance the amount of enzyme giving specific signal versus that giving background signal. The most effective way to control this is by optimizing the enzyme conjugate, antibodies, and blocking solution.
3) Poor standard curve linearity
If the standard curve has a poor linearity, then samples must fall within a tight concentration range to be deemed accurate.
If the curve has good linearity but poor variation between replicates (i.e., standard error), there might be a technical problem such as inconsistent pipetting between samples or individual users.
All samples and standards should be measured at least in duplicate or triplicate to mitigate this issue.
4) Excessively high signal in ELISA
Insufficient washing. To avoid this issue, use the appropriate washing procedure, e.g., at the end of each washing step, invert the plate on absorbent tissue and allow to completely drain, tapping forcefully if necessary to remove any residual fluid.
Incubation time is too long. Excessive incubation time is also a reason for overly high signal in ELISA be sure to follow recommended incubation times.
Contamination. Avoid this issue using plate sealers, and use a fresh sealer each time the plate is opened. This will prevent wells from contaminating each other.