Troubleshooting

Little or no signal may be due to: (1) the primary antibody concentration, (2) the cell collection method, or (3) the flow cytometry assay itself.

• Please check that your secondary antibody recognizes the primary antibody: most of our antibodies are generated in rabbits.
• If you have previously used your secondary antibody with successful results, we suggest trying longer incubation times with the primary antibody (up to 2 hours at 4°C) in a smaller reaction volume.
• Your primary antibody concentration may be too low. Try using more primary antibody or set up a series of dilutions to determine the optimal concentration for your system.
• If you are trying to stain an intracellular protein, you’ll need to permeabilize your cells first.
• If you are attempting to detect cell surface proteins, they may have become internalized. Trypsin can induce the internalization of cell surface proteins so you may need to alter the cell detachment method. You can reduce the internalization of cell surface proteins by adding NaN₃ or by keeping your assay reagents on ice.

High background levels are most likely due to too much primary antibody concentration or insufficient blocking. Both of these cases can independently lead to a high signal in what should be negative cell populations.

• If you are using direct flow cytometry, use a suitable conjugated IgG isotype control.
• If you are using indirect flow cytometry, use your secondary antibody alone as a negative control.
• Too much antibody will produce a lot of background. Test different antibody dilutions to determine the optimal concentration.
• Your antibody may be binding off-targets such as the Fc receptors. You can try an Fc receptor blocking reagent to minimize the off-target binding.
• There may be doublets in your cell population, i.e., dividing cells. Try adding EDTA to your buffer or filtering the cells through a 30 µm filter.
• It’s possible that the flow cytometer equipment settings may need adjusting: if the offset is too low, or the gain too high, you will generate background signal.
• If you are not using a conjugated primary antibody, you may need to simply increase the wash times, or add extra wash steps to your protocol.
• When running multiple fluorochromes, spillover can occur. Ensure that you have used a multicolor panel building to confirm that you have a workable combination of fluorochromes.
• When choosing the fluorochromes, the brightest fluorochrome for the target with the lowest expression or density should be selected. Conversely, choose the dimmest fluorochrome for those targets with the highest expression or density.
• If you have high side scatter background it could be due to bacterial contamination, which will autofluoresce and give high event rates.

If your flow cytometer cannot consistently distinguish between individual cells, your event rate will appear abnormal. Remember, you may need to obtain as many as 10⁷ events for significant detection.

• If the event rate is too low, ensure that the cells are properly mixed and that the population is between 1 x 10⁵ and 1 x 10⁶ cells/ml.
• If the event rate is too low, your cells could be clumping together. Make sure you sieve the cells before they are acquired and sorted to remove any debris.

All of our antibodies undergo strict QC analysis by western blotting. We purify each lot with an affinity column, and we use the same protocol and conditions throughout the analysis. Also, we only release our antibodies after we obtain satisfactory results in comparison with the previous lot. If you do not obtain a signal, there could be a fundamental issue with your antibody or technique.

• You may not have used enough primary or secondary antibody. Please follow the recommended antibody dilutions or test different antibody dilutions to determine the optimal antibody concentrations.
• If you’re using a fluorescent detection system, you may not have kept your conjugated primary or secondary antibody in the dark. Ensure that these antibodies are not exposed to light whenever possible.
• There’s the possibility that your primary and secondary antibodies do not work together. Ensure that the secondary antibody was raised against the animal that the primary antibody was raised in.
• The NaN₃, present in the antibody solution, can sometimes cause problems.
• Your fixation method could be damaging the epitope and preventing the primary antibody from recognizing it. Either reduce the fixation time or try an antigen retrieval method.
• Your protein of interest may not be present or may be present at very low levels, in the sample that you’re testing. Alternatively, the primary antibody may not recognize the protein of interest in the species that you’re testing. If you are certain the protein is expressed, you can try an enrichment step to improve the signal. You should also check the antibody datasheet to ensure that it cross-reacts with the species that you’re testing.

High background levels may be due to a sub-optimal primary antibody concentration, an insufficient blocking step, or there could be non-specific binding between the antibodies and the blocking reagent.

• Titrate the primary antibody to obtain the optimal concentration. Check the datasheet for the optimal dilution, but we recommend 1:100 as a starting point for most primary antibodies. For antibodies directly conjugated to a fluorophore, we suggest a 1:50-1:60 dilution.
• Try adding 0.1%–0.5% of Tween-20 to the antibody solution and washing buffer since this strengthens the signal and reduces non-specific staining. Tween-20 is more gentle than other detergents such as Triton X-100, which can damage the membrane during permeabilization.
• In some cases, the biotin-avidin amplification can cause high background and there is endogenous avidin binding ability in several tissues. Thus, when we use a biotin-conjugated secondary antibody and then add extravidin or streptavidin conjugated to peroxidase or a fluorophore, it may be bound not only to the biotin from the secondary antibody but also to binding sites in the tissue. To overcome this problem, saturate endogenous binding sites before exposure to the primary antibody by incubation of the tissue sections in a solution containing non-conjugated streptavidin. We use a kit from Vector Laboratories (US): kit SP-2002.
  • These steps are for section after antigen retrieval treatments (if required) and after 2 x 5-minute rinses in IHC-PBS.
  • Pre-saturate endogenous avidin binding ability: Add 4 drops from the “streptavidin” bottle to 5 ml of IHC-PBS.
  • Incubate sections in this solution 30 minutes at room temperature.
  • Rinse 2 x 5 minutes in IHC-PBS.
  • Saturate residual biotin binding potential: Add 4 drops from the “biotin” bottle to 5 ml of PBS. Incubate sections in this solution 30 min at room temperature.
  • Rinse 2 x 5 minutes in IHC-PBS.
  • Start IHC and transfer sections to the primary antibody.
• Your sample incubation temperature may be too high. Incubate the tissue sections at 4°C.
• Excessive fixation times can affect the epitope and cause high background levels. Try a fixation protocol with a lower exposure time, lower temperature, and/or reduced concentration of fixative. In general, we incubate the sample for 24–72 hours in 4% paraformaldehyde (PFA). When the samples consist of bloody organs, such as spleen and liver, we recommend you replace the PFA after 24 hours.
• If you grew your cells on a coated chamber slide, the antibody may stick to the surface. In this case, try using another coating solution (e.g., polyethylene glycol, polyethyleneimine, etc.) since it may affect the background levels.

This is due to endogenous peroxidase activity. Try endogenous peroxidase quenching with hydrogen peroxide. For more details, check out our IHC protocol for paraffin embedded sections.

This is likely due to damaged tissue from sectioning or other similar procedures.

• Aggregates may form in the antibody solution following reconstitution and especially after thawing of the reconstituted antibody solution. Thus, we recommend centrifuging all of the antibody preparations before use (10000 x g for 5 min).
• Sectioning with a dull blade can cause folding or air bubbles. Moreover, cutting sections too thick can make them difficult to resolve. Ensure your section equipment is well-maintained, and your protocol is optimized.

We find that floating sections are stained more effectively compared to sections adhered to slides. When sections are adhered to slides, there are several potential problems since the access and penetration of the antibody beyond the surface of the sections is limited. You can overcome this to some degree by increasing the Triton X-100 content in the antibody solution. However, we don’t recommend this in many cases where the antibody targets an extracellular epitope because Triton X-100 is a relatively harsh detergent and too much can deteriorate the antigens.

All of our antibodies undergo strict QC analysis by western blotting. We purify each lot with an affinity column, and we use the same protocol and conditions throughout the analysis. Also, we only release our antibodies after we obtain satisfactory results in comparison with the previous lot. If you do not obtain a signal, there could be a fundamental issue with your antibody or technique.

• You may not have used enough primary or secondary antibody. Please follow the recommended dilutions for the antibodies or test different antibody dilutions to determine the optimal antibody concentrations.
• There’s the possibility that your primary and secondary antibodies do not work together. Ensure that the secondary antibody was raised against the animal that the primary antibody was raised in.
• The NaN₃, present in the washing buffer and secondary antibody solution, can sometimes cause problems.
• Your protein of interest may not be present or could be present at very low levels in the sample. Alternatively, the primary antibody may not recognize the protein from the species that you’re testing. If you are certain the protein is expressed, you can try an enrichment step to improve the signal. You should also check the antibody datasheet to ensure that the antibody cross-reacts with the species you’re using.
• Try loading more protein into your gels or use an alternative protein extraction method for sample preparation. Remember, the optimal sample preparation method for your target protein may not necessarily be identical to the one used for a control housekeeping protein (e.g., GAPDH).

The expected molecular weight (MW) listed in the product datasheet is based solely on the size of the target protein’s amino acid sequence. It’s important to remember that many factors can affect the banding pattern of your western blot including: (1) the existence of a splice variant, (2) the quality of the loaded sample, (3) the protein extraction method, and (4) the protein transfer conditions. To achieve accurate results, you may need to systematically adjust the protein extraction method or the protein transfer conditions.

• If the band’s MW is below the expected MW it could be due to a splice variant with a slightly different MW, or the target protein may have undergone proteolytic cleavage, generating a lower MW band. Changing your protein extraction method or transfer conditions may resolve this issue. You can also try the following steps:
  • Heat the samples at 70°C for 10 min (instead of 100°C for 5 min) as this reduces potential degradation.
  • Increase the transfer time. This will increase the transfer efficiency of high molecular weight (HMW) proteins. We recommend 100 mA for 20 h at 4°C.
• If the band’s MW is above the expected MW, it could be due to post-translational modifications (e.g., glycosylation or phosphorylation) or the existence of a different splice variant.
• In either event, you should use a blocking peptide as a negative control. This peptide is the antigen used for immunization and specifically blocks the antibody. So, in the presence of the blocking peptide, specific staining will disappear while non-specific staining will remain. You should use the blocking peptide at the ratio indicated in the datasheet, or 1 µg of peptide per µg of antibody.

High background levels may be due to the following: (1) a sub-optimal primary antibody concentration, (2) the type of membrane, (3) the blocking conditions, or (4) non-specific binding between the antibodies and the blocking reagent.

• Titrate the primary antibody to obtain the optimal concentration. Check the datasheet for the optimal dilution, but we recommend 1:200 in most cases.
• You may consider changing to a nitrocellulose membrane. Compared with polyvinylidene difluoride (PVDF) membranes, a nitrocellulose membrane tends to give you less background noise but does lack some of the detection sensitivity of a PVDF membrane.
• Also, ensure that the membrane doesn’t dry out during the western blot.
• If you haven’t blocked for long enough, you can end up with a lot of background in your WB. We recommend an hour with 1–3% bovine serum albumin (BSA), but overnight at 4°C can also work.
• Try adding 0.1%–0.5% of Tween-20 to the primary antibody solution and washing buffer since this strengthens the signal and reduces non-specific staining.

Usually, too many bands indicate that the sample preparation method was not optimal and/or that the target protein was degraded during sample preparation.

• Multi-transmembrane proteins like such as calcium channels are usually vulnerable to degradation during sample preparation. The optimal sample preparation method can vary depending on the tissue (e.g., brain, adrenal gland, etc.) and/or species (e.g., mouse, human, etc.). For this reason, we normally recommend running different samples prepared by different protocols in parallel, to establish the optimal sample preparation method. Remember, the optimal sample preparation method for your target protein may not necessarily be identical to the one used for a control housekeeping protein (e.g., GAPDH).
• If you are analyzing HMW proteins, you may need to adjust the SDS-PAGE conditions and the transfer conditions.
• Additional bands may indicate that there is a splice variant or a truncated version of your target protein.
• You can try a more diluted primary antibody concentration and/or add up to 0.5% Tween-20 to the primary antibody solution.

To make sure you get optimal blocking of the primary antibody, incubate the antibody, in parallel, with and without the antigen (the antibody/antigen ratio is available in the certificate of analysis delivered with the antibody) in a small volume (500 µl of 1% BSA in PBS) for 1 hour at room temperature with rotation. After the incubation, dilute each vial to the appropriate working concentration in the desired buffer and apply the contents of each vial to each membrane for parallel experiments.

• We recommend that you use the blocking peptide at the ratio specified in the datasheet, or 1 µg of peptide per µg of antibody.
• If you’re using a fusion protein to block the antibody, be aware that fusion proteins are sometimes more delicate to handle than peptides. If the fusion protein was correctly reconstituted (by adding 100 µl of PBS) and handled (avoidance of repeat freezing and thawing), a more diluted antibody together with an increased amount of fusion protein may be necessary for complete blocking of the antibody. We normally recommend adding 3 µg of fusion protein for each µg of antibody, but sometimes you need a larger ratio depending on the sample.

These curved bands (sometimes called “smiling bands”) are likely due to excessively high voltage, which causes excessively rapid migration of the proteins. Smiling bands can also occur when the temperature is too high.

• Reduce the voltage and run the gel in a cooler room.

To help identify the cause, consider the following:

1. Verify that the target ion channel subtype is consistent with the reported pharmacological profile of the toxin.
2. Evaluate a range of toxin concentrations and incubation times appropriate for your experimental system.
3. Compare assay conditions with those reported in the literature, including cell type, recording conditions, and assay temperature.
4. Ensure the vial is centrifuged before opening to collect any lyophilized material on the walls or cap, and is reconstituted according to the product webpage and datasheet instructions.
5. Confirm that the solvent concentration (e.g., DMSO) used for reconstitution does not inherently affect or suppress the target ion channel current.

To help identify the cause, consider the following:

1. Confirm that the target channel is expressed and produces a stable, measurable current.
2. Verify that the toxin concentration and incubation time are appropriate for the assay.
3. Ensure the vial is centrifuged before opening to collect any lyophilized material on the walls or cap, and is reconstituted according to the product webpage and datasheet instructions.
4. Use fresh toxin aliquots and avoid repeated freeze-thaw cycles.
5. Confirm recording quality, solution composition, voltage protocol, and assay temperature.
6. Include an appropriate positive control, when available.

A low or absent signal in FACS staining with conjugated toxins may result from suboptimal staining conditions, insufficient toxin concentration, or improper incubation parameters. The following steps may help improve the staining intensity:

1. Optimize the toxin concentration
   Test a range of toxin concentrations to determine the optimal level for your experiment. Concentrations that are too low may fail to produce detectable staining. A starting titration range of 0.1-1 µM is generally recommended, although the optimal concentration may vary depending on the target expression level and cell type.
   
   
2. Adjust the incubation temperature
   Incubating cells with the conjugated toxin at 37°C are highly recommended, as it often results in stronger and more consistent staining compared to incubation at 4°C. Higher temperatures can improve toxin-target interactions and may facilitate better accessibility to the target proteins on the cell surface.
   
   
3. Ensure sufficient incubation time
   In some cases, extending the incubation time (e.g., 30-60 minutes) can improve binding and increase signal intensity, especially when target expression levels are low.
   
   
4. Verify target expression with proper controls
   Confirm that the cells used in the experiment express the toxin’s target. Include positive controls, such as transfected cells or stable cell lines expressing the target. These controls can help determine whether the staining protocol is working correctly.
   
   
5. Preserve cell surface protein integrity
   Make sure that the cells are viable and properly handled prior to staining. Damage to the cell membrane, such as permeabilization and fixation, or improper preparation can affect the native conformation of targets and reduce toxin binding.

1. Optimize toxin concentration
   Ensure that the toxin is used within an appropriate concentration range. Using concentrations that are too high can increase background and lead to non-specific binding. It is recommended to test a titration range (e.g., 0.1-1 µM as a starting point) to identify the optimal concentration for your experiment.
   
   
2. Use appropriate controls
   Include proper controls to confirm staining specificity. For example:
   a. Positive control: cells that express the toxin’s target (e.g., transfected cells or stable cell lines).
   b. Negative control: cells lacking the target, such as KO cells, when available.
   
   
3. Avoid cell or tissue permeabilization
   Do not permeabilize cells or tissues when staining with venom toxins. Permeabilization allows toxins to enter the cell, which may increase non-specific binding to intracellular components. Importantly, venom toxins typically recognize specific extracellular structural elements of their target proteins, such as pore-forming regions or voltage-sensing domains. Therefore, preserving the native conformation of target proteins is essential for accurate and reliable staining.
   
   
4. Validate specificity with complementary markers
   When performing multiplex staining, it is useful to validate toxin binding using Alomone’s antibodies that recognize extracellular epitopes on the same protein target. This helps confirm that the observed staining corresponds to the intended target.

1. Optimize the toxin concentration
   Test a range of toxin concentrations to determine the optimal level for your experiment. Concentrations that are too low may fail to produce detectable staining. A starting titration range of 0.1-1 µM is generally recommended, although the optimal concentration may vary depending on the target expression level and cell type.
   
   
2. Adjust the incubation temperature
   Incubating live cells with the conjugated toxin at 37°C is highly recommended, as it results in stronger and more consistent staining compared to incubation at 4°C. Higher temperatures can improve toxin-target interactions and may facilitate better accessibility to the target proteins on the cell surface.
   
   
3. Ensure sufficient incubation time
   Incubation time can vary depending on several factors, including the potency of the toxin and the level of target expression in the cells being analyzed. Highly potent toxins or cells with high target expression may require shorter incubation periods, whereas lower toxin activity, always ensuring you are working within the recommended effective concentration range as specified in the product specifications, or reduced target expression may necessitate longer exposure times to achieve adequate staining.
   
   For this reason, it is recommended to empirically optimize the incubation conditions for each experimental setup. Performing preliminary calibration experiments to determine the appropriate incubation duration will help maximize staining specificity and signal quality while minimizing non-specific background. Careful optimization of incubation timing is therefore essential for obtaining reliable and reproducible toxin staining results.
   
   
4. Prevent cell detachment
   Detachment of adherent cells from their substrate or from neighboring cells can significantly alter target protein activity due to associated mechanical, biochemical, and structural changes in the cells. These changes may affect the physiological state of the cells and influence experimental outcomes.
   
   Therefore, when performing washing steps, cells should be handled very gently to avoid applying excessive force that could cause cell detachment. Careful pipetting and minimal disturbance of the cell layer are recommended to maintain normal cellular conditions.
   
   For live-cell imaging protocols, it is also recommended to use phosphate-buffered saline (PBS) containing calcium and magnesium during washing steps, as these ions help maintain cell-cell and cell-substrate adhesion and reduce the likelihood of cell detachment.
   
   
5. Avoid photobleaching and cytotoxicity
   Avoid using high laser power during image acquisition, as excessive laser intensity can cause rapid photobleaching of the fluorophores and may also induce phototoxic effects that compromise cell viability. These effects can reduce signal quality over time and negatively impact the reliability of live-cell imaging experiments.
   
   To preserve the fluorescence signal and maintain high viability of the cells, it is recommended to use the lowest laser power that still provides an adequate signal-to-noise ratio. Optimizing detector gain, exposure time, and scanning settings can further help improve image quality while minimizing photodamage.
   
   In addition, utilizing advanced imaging systems can significantly enhance signal detection and resolution. This also enables super-resolution imaging with improved sensitivity, allowing researchers to capture high-quality images at lower laser intensities while resolving finer structural details.

To help identify the cause, consider the following:

1. Verify that the target cells express the relevant receptor.
2. Optimize growth factor concentration and treatment duration.
3. Ensure the vial is centrifuged before opening to collect any lyophilized material on the walls or cap, and is reconstituted according to the product webpage and datasheet instructions.
4. Standardize cell culture conditions, including passage number, cell density, and serum conditions.
5. Consider serum reduction or serum starvation where appropriate for the assay.
6. Confirm assay performance using an appropriate positive control and, when available, a known responsive cell model.
7. Aliquot the reconstituted protein upon opening to avoid repeated freeze-thaw cycles, which can cause rapid degradation.

To help identify the cause, consider the following:

1. Verify that the assay endpoint is being measured at an appropriate time point.
2. Confirm that the target cells express the relevant receptor at sufficient levels.
3. Consider that different cell types may exhibit different sensitivities to the same growth factor.
4. Evaluate whether serum components may interfere with or mask the biological response.
5. Compare your protocol with the activity assay described in the product datasheet.
6. Confirm assay sensitivity by including an appropriate positive control.
7. Consider evaluating additional downstream signaling endpoints or biomarkers.
8. Ensure the vial is centrifuged before opening to collect any lyophilized material on the walls or cap, and is reconstituted according to the product webpage and datasheet instructions.

A low or absent signal in FACS staining with conjugated neurotrophins may result from suboptimal staining conditions, insufficient neurotrophin concentration, or improper incubation parameters. The following steps may help improve staining intensity:

1. Optimize the neurotrophin concentration
   Test a range of neurotrophin concentrations to determine the optimal level for your experiment. Concentrations that are too low may fail to produce detectable staining. A starting titration range of 10-100 nM is generally recommended, although the optimal concentration may vary depending on the target expression level and cell type.
   
   
2. Adjust the incubation temperature
   Incubating cells with the conjugated neurotrophin at 37°C are highly recommended, as it often results in stronger and more consistent staining compared to incubation at 4°C. Higher temperatures can improve neurotrophin-receptor interactions and may facilitate better accessibility to the target proteins on the cell surface.
   
   
3. Ensure sufficient incubation time
   In some cases, extending the incubation time (e.g., 30-60 minutes) can improve binding and increase signal intensity, especially when receptor expression levels are low.
   
   
4. Verify receptor expression with proper controls
   Confirm that the cells used in the experiment express the neurotrophin-specific receptor. Include positive controls, such as transfected cells or stable cell lines expressing the target receptor, which may help determine whether the staining protocol is working correctly.
   
   
5. Preserve cell surface protein integrity
   Make sure that the cells are viable and properly handled prior to staining. Damage to the cell membrane or improper preparation can lead to receptor degradation and reduce binding affinity.

Several factors may explain why cells do not respond to neurotrophin stimulation:

1. Lack of receptor expression
   Cells must express the appropriate neurotrophin receptors (e.g., TrkA, TrkB, TrkC, or p75ᴺᵀᴿ) to respond to treatment. If these receptors are not present or are expressed at very low levels, the cells may not respond to the neurotrophin.
   
   
2. Suboptimal neurotrophin concentration
   The concentration used may be too low to induce a measurable biological response. Optimizing the concentration range and performing dose-response experiments can help determine the appropriate working concentration.
   
   
3. Cell viability and culture conditions
   Cells that are stressed, over-confluent, or not maintained under optimal culture conditions may show reduced responsiveness to neurotrophins.
   
   
4. Inappropriate experimental timing
   Neurotrophin-induced responses such as signaling activation, survival, or differentiation may occur at different time points. It may be necessary to optimize the duration of treatment depending on the biological endpoint being measured.
   
   
5. Assay sensitivity
   The detection method used to measure the response (e.g., survival assay, phosphorylation assay, or gene expression analysis) may not be sensitive enough to detect subtle changes.
   
   
6. Neurotrophin stability or handling
   Improper storage, repeated freeze-thaw cycles, or prolonged exposure at room temperature (RT) can reduce neurotrophin activity. Always follow the recommended storage and handling conditions.
   
   
7. Cell type-specific responses
   Different cell types respond differently to neurotrophins. Some cells may require additional co-factors or specific culture conditions to exhibit a detectable response.

1. Optimize neurotrophin concentration
   Ensure that the neurotrophin is used within an appropriate concentration range. Using concentrations that are too high can increase background and lead to non-specific binding. It is recommended to test a titration range (e.g., 10-100 nM as a starting point) to identify the optimal concentration for your experiment.
   
   
2. Use appropriate controls
   Include proper controls to confirm staining specificity. For example:
   a. Positive control: cells that express the specific neurotrophin receptor (e.g., transfected cells or stable cell lines).
   b. Negative control: cells lacking the target receptor, such as knockout (KO) cells, when available.
   
   
3. Validate specificity with complementary markers
   When performing multiplex staining, it is useful to validate the specificity of neurotrophin binding using Alomone`s antibodies that recognize the receptors of different neurotrophins. This helps confirm that the observed staining corresponds to the intended target.

To help identify the cause, consider the following:

1. Confirm receptor expression and functional coupling in the selected cell system.
2. Optimize ligand concentration and stimulation time.
3. Ensure the vial is centrifuged before opening to collect any lyophilized material on the walls or cap, and is reconstituted according to the product webpage and datasheet instructions.
4. Confirm that the solvent concentration (e.g., DMSO) used to dissolve the ligand does not inherently induce a calcium response, affect cell viability, or interfere with assay detection.
5. Confirm assay performance using an appropriate positive control.
6. Use healthy, low-passage cells at consistent density.
7. Confirm proper dye loading and assay buffer conditions.
8. Minimize repeated stimulation, which may cause receptor desensitization.

To help identify the cause, consider the following:

1. Maintain consistent cell passage number, cell density, and culture conditions.
2. Standardize dye loading concentration, time, temperature, and washing steps.
3. Standardize ligand preparation, dilution, and addition timing across experiments.
4. Avoid repeated receptor stimulation before the assay.
5. Confirm that assay temperature, plate reader settings, and acquisition timing are consistent between experiments.
6. Include internal controls on each plate.

To help identify the cause, consider the following:

1. Verify receptor expression and membrane localization in the selected cell system.
2. Confirm that ligand concentration and incubation conditions are appropriate.
3. Ensure the vial is centrifuged before opening to collect any lyophilized material on the walls or cap, and is reconstituted according to the product webpage and datasheet instructions.
4. Verify that the solvent concentration (e.g., DMSO) used in the final assay buffer does not disrupt ligand-receptor binding or interfere with the detection system.
5. Optimize incubation time, temperature, buffer composition, and washing conditions.
6. Include appropriate positive and negative controls, when available.
7. For competition assays, confirm that the competitor concentration range is appropriate.

To help identify the cause, consider the following:

1. Compare cell type, receptor expression system, receptor species, and receptor density.
2. Compare ligand concentration, incubation time, temperature, buffer composition, and washing conditions.
3. Confirm whether the published study used live cells, membranes, recombinant cells, or native tissue.
4. Check whether the assay endpoint and data analysis method are equivalent.
5. Include appropriate reference controls to evaluate assay performance.
6. Minor differences in assay design, receptor expression levels, or experimental conditions can significantly affect binding values and apparent potency.
7. Ensure the vial is centrifuged before opening to collect any lyophilized material on the walls or cap, and is reconstituted according to the product webpage and datasheet instructions.