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Base Cat #A78291

Symmetric DiMethyl-Histone H3-R8 Polyclonal AntibodyChIP Grade

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Recognizes Symmetric Dimethylated Histone H3-R8 as a histone methylation mark in histone modification and chromatin regulation.

Application: ChIPELISAWB
Clonality: Polyclonal
Conjugate: Unconjugated
Host: Rabbit
Isotype: IgG
Purification: Affinity Purified
Reactivity: Broad Range, Human, Mouse, Rat
100% Guarantee: 6 months
Application Guarantee for AntibodiesRecombinant Rabbit Monoclonal Abs
Catalog No.SizePriceQty
A78291-100100 µl $368.00 
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Product Overview

Background
Histones are basic nuclear proteins that are responsible for the nucleosome structure of the chromosomal fiber in eukaryotes. This structure consists of approximately 146 bp of DNA wrapped around a nucleosome, an octamer composed of pairs of each of the four core histones (H2A, H2B, H3, and H4). The chromatin fiber is further compacted through the interaction of a linker histone, H1, with the DNA between the nucleosomes to form higher order chromatin structures. This gene is intronless and encodes a replication-dependent histone that is a member of the histone H3 family. Transcripts from this gene lack polyA tails; instead, they contain a palindromic termination element. This gene is found in the large histone gene cluster on chromosome 6p22-p21. 3.

Description
Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody. Unconjugated. Raised in: Rabbit.

Formulation
Buffer: PBS containing 50% glycerol, preserved with proclin300 or sodium azide, pH 7.3.

Specificity
Human, Mouse, Rat, Other (Broad Range)

Isotype
IgG

Uniprot ID
Q16695/P68431

Purification
Affinity Purified

Immunogen
Synthetic peptide. Please contact EpigenTek for more information.

Storage
Store at -20°C. Avoid freeze / thaw cycles.

Alternative Names
H3/A; H3C2; H3C3; H3C4; H3C6; H3C7; H3C8; H3FA; H3C10; H3C11; H3C12; HIST1H3A; Symmetric DiMethyl-Histone H3-R8

Application
WB, ELISA, ChIP; Recommended dilution: WB, 1:200; ELISA, 1:1000 - 1:2000

Datasheet & SDS

PDF
Datasheet/User Guide: A78291 If the above file is not available online, you can request a copy by contacting us.

Applications

Suggested protocols for Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody should be self-optimized according to the user's experiment.

Choose Your Experiment Type

Use this Western blot protocol guide to choose conditions for sample preparation, SDS-PAGE gel selection, protein transfer, membrane blocking, antibody incubation, detection, controls, and troubleshooting. Select the experiment type that best matches your sample or target, including routine lysate, recombinant protein, tagged protein, low-abundance target, small protein or histone, large protein, membrane protein, phospho/PTM target, IP-Western or pulldown eluate, or secreted protein.

Routine cell or tissue lysate Western blot protocol

For most 20-100 kDa targets in whole-cell lysate or tissue lysate.

  • Use this workflow when: You are detecting a typical 20-100 kDa target in whole-cell lysate, nuclear extract, cytoplasmic extract, or tissue lysate.
  • Optimize first: Sample quality, protein normalization, transfer consistency, antibody dilution, blocking buffer, and exposure time.

Before you start

  • Confirm the expected molecular weight and the antibody host species before starting.
  • Prepare fresh protease inhibitors in cold lysis buffer. For nuclear protein lysates, consider EpiQuik Nuclear Extraction Kit (OP-0002); for histone protein lysates, consider EpiQuik Total Histone Extraction Kit (OP-0006).
  • Plan 10-30 ug total protein per lane as the starting load.
  • Include a positive control, negative control when available, and a validated loading control or total protein stain.

Protocol

1Prepare and clarify the sample

  1. For cells, collect cells directly and wash with cold PBS when appropriate. For tissues, cut and homogenize tissue pieces, then collect and wash the resulting cell pellet.
  2. Lyse on ice in your own cold lysis buffer containing fresh protease inhibitors, or use the reference lysis buffer preparation below. For nuclear protein lysates, consider EpiQuik Nuclear Extraction Kit (OP-0002); for histone protein lysates, consider EpiQuik Total Histone Extraction Kit (OP-0006).
  3. Incubate on ice for 10-30 minutes with occasional mixing.
  4. Clarify at 12,000-16,000 x g for 10-20 minutes at 4°C.
  5. Transfer only the clear supernatant to a fresh tube. If lysate is viscous, shear DNA before loading.

2Quantify, normalize, and denature

  1. Measure protein concentration before adding reducing sample buffer.
  2. Normalize all samples to the same concentration.
  3. Add 4X Laemmli sample buffer to 1X final concentration.
  4. Heat at 95°C for 5 minutes, then briefly centrifuge before loading.

3Run SDS-PAGE

  1. Use a 10-12% gel or 4-12% gradient gel for most routine targets.
  2. Load ladder, positive control, negative control, and test samples.
  3. Run the stacking gel at 60-80 V, then the resolving gel at 100-150 V.
  4. Stop when the dye front approaches the bottom or the target range is separated.

4Transfer to membrane

  1. Use 0.45 um PVDF or nitrocellulose for routine targets.
  2. For wet transfer, start around 100 V for 60-90 minutes at 4°C, or use validated semi-dry settings.
  3. Stain the membrane with Ponceau S before blocking to confirm transfer and lane consistency.
  4. Correct uneven transfer before changing antibody conditions.

5Block, probe, detect, and normalize

  1. Block with 5% non-fat dry milk in TBST unless the antibody datasheet recommends another blocker.
  2. Incubate Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody using the dilution, incubation time, and buffer recommended in the antibody datasheet or supplier specifications.
  3. Wash 3 x 5-10 minutes, incubate secondary antibody, then wash again.
  4. Capture non-saturated exposures and normalize to total protein or a validated loading control.

Controls to include

  • Positive lysate or purified protein that is known to contain the target.
  • Negative lysate, knockout, knockdown, untreated, or low-expression sample when available.
  • Loading control such as GAPDH, beta-actin, tubulin, or a total protein stain.
  • Short and long exposures to avoid relying on a saturated image.

Purified or recombinant protein Western blot protocol

For purified target protein, recombinant controls, standards, or defined protein preparations.

  • Use this workflow when: You are confirming expression, purity, identity, tag detection, or recovery of a purified or overexpressed recombinant protein.
  • Optimize first: Protein loading amount, tag or target antibody choice, gel percentage, transfer time, and exposure range.

Before you start

  • Use nanogram-range loading instead of total lysate loading ranges.
  • Prepare a dilution series so signal intensity can be interpreted without saturation.
  • Confirm the final expected size, including tag, linker, fusion partner, or cleavage product.
  • Include unrelated recombinant protein, buffer-only control, or tag-only control when relevant.

Protocol

1Prepare a dilution series

  1. Dilute the protein in compatible buffer before adding sample buffer.
  2. Start with 1-50 ng per lane for most purified proteins.
  3. Include at least three amounts, such as low, medium, and high load.
  4. Use a buffer-only lane if the storage buffer contains reducing agent, salt, glycerol, detergent, or carrier protein.

2Denature without overloading

  1. Add 4X sample buffer to 1X final concentration.
  2. Heat at 95°C for 5 minutes unless the protein aggregates.
  3. If precipitation appears after heating, test 37-70°C for 10-30 minutes.
  4. Briefly centrifuge before loading.

3Run SDS-PAGE by final protein size

  1. Choose gel percentage by the final expected recombinant protein size.
  2. Use a gradient gel if expected contaminants or fragments span a wide molecular weight range.
  3. Run the gel until the target and any expected fragments are separated.
  4. Do not interpret an overloaded purified lane as antibody specificity.

4Transfer

  1. Use 0.45 um membrane for most recombinant proteins above about 20 kDa.
  2. Use 0.2/0.22 um membrane for very small proteins or peptides.
  3. Confirm transfer with Ponceau S or another reversible stain.
  4. If signal is too intense, reduce protein load before changing transfer conditions.

5Probe and interpret

  1. Start with datasheet antibody dilution or a small antibody titration.
  2. Capture exposure times that keep the dilution series within the non-saturated range.
  3. Use unrelated recombinant protein or tag-only protein to check non-specific recognition.
  4. Use lysate validation separately if the antibody will be used on complex samples.

Controls to include

  • Purified target protein as the main positive control.
  • Unrelated recombinant protein to check non-specific recognition.
  • Tag-only control if the detection antibody recognizes a tag.
  • Buffer-only lane to identify artifacts from storage buffer.

Tagged protein expression Western blot protocol

For FLAG, HA, His, Myc, GFP, or other tagged proteins expressed in cells or host lysates.

  • Use this workflow when: You are detecting HA, FLAG, His, Myc, GFP, or another tagged construct in transfected, transduced, or engineered samples.
  • Optimize first: Tag accessibility, expression level, lysis conditions, positive and mock-transfected controls, and antibody dilution.

Before you start

  • Calculate the final expected molecular weight, including tag, linker, fusion partner, and cleavage site.
  • Prepare matched samples such as transfected vs mock, induced vs uninduced, or expression vector vs empty vector.
  • Select lysis conditions based on tagged protein localization. For tagged nuclear proteins, consider EpiQuik Nuclear Extraction Kit (OP-0002); for tagged histone proteins, consider EpiQuik Total Histone Extraction Kit (OP-0006).
  • Use anti-tag antibody to confirm expression and target-specific antibody to confirm identity when possible.
  • Plan lower lysate loads if expression is very strong.

Protocol

1Prepare matched expression lysates

  1. Collect test and control samples under the same lysis conditions.
  2. Lyse on ice in your own cold lysis buffer containing fresh protease inhibitors, or use the reference lysis buffer preparation below. For nuclear protein lysates, consider EpiQuik Nuclear Extraction Kit (OP-0002); for histone protein lysates, consider EpiQuik Total Histone Extraction Kit (OP-0006).
  3. Centrifuge lysates at 12,000-16,000 x g for 10-20 minutes at 4°C.
  4. Start with 10-30 ug total protein per lane, or less if overexpression is strong.
  5. Keep induced/uninduced or transfected/mock samples paired on the same gel.

2Normalize and denature

  1. Quantify protein and normalize all lysates.
  2. Add reducing sample buffer to 1X final concentration.
  3. Heat at 95°C for 5 minutes unless the tagged protein aggregates.
  4. If the protein smears or remains in the well, test lower-temperature denaturation.

3Run gel by final tagged size

  1. Choose gel percentage based on the final construct size, not the native protein alone.
  2. Use 10-12% gel for many 25-100 kDa constructs, higher percentage for small tags, and lower percentage for large fusions.
  3. Run the gel long enough to separate full-length target from degradation products.
  4. Use an appropriate molecular weight ladder covering the full construct size.

4Transfer

  1. Transfer based on final target size.
  2. Use 0.45 um membrane for most tagged proteins above about 20 kDa.
  3. Check transfer before blocking.
  4. If the target is a large fusion protein, use large-protein transfer settings.

5Probe with tag and target logic

  1. Probe with anti-tag antibody to confirm expression.
  2. Probe with target-specific antibody when identity needs confirmation.
  3. Compare signal against mock, empty-vector, or uninduced control.
  4. If anti-tag gives many bands, reduce load and check for degradation.

Controls to include

  • Mock or untransfected lysate to identify host-cell background.
  • Empty-vector lysate to control for tag-related signal.
  • Induced and uninduced samples for inducible expression systems.
  • Target-specific antibody confirmation when anti-tag signal alone is not enough.

Low-abundance endogenous target Western blot protocol

For weak endogenous targets, rare proteins, inducible proteins, or targets that are difficult to detect in total lysate.

  • Use this workflow when: The target is weak, rare, inducible, tissue-restricted, or expected to produce a low signal in routine lysate.
  • Optimize first: Positive control choice, sample enrichment, protein load, Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody incubation, wash stringency, and exposure time.

Before you start

  • Identify a strong positive control before running unknown samples.
  • Consider enrichment, IP, fractionation, nuclear extraction, or membrane enrichment instead of simply loading more lysate.
  • Use fresh inhibitors and keep samples cold.
  • Plan overnight Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody incubation and sensitive detection.

Protocol

1Prepare or enrich the target

  1. Use total lysate only if the target is expected to be detectable without enrichment.
  2. For nuclear, membrane, chromatin-associated, or low-copy targets, enrich the appropriate fraction first. For nuclear protein enrichment, consider EpiQuik Nuclear Extraction Kit (OP-0002).
  3. Start with 20-40 ug total lysate only if lane quality remains acceptable.
  4. Include the positive control on the same blot.

2Normalize and denature

  1. Quantify protein or normalize enriched fraction by a consistent basis.
  2. Add reducing sample buffer and heat at 95°C for 5 minutes unless the target requires another condition.
  3. Avoid overloading total lysate because it can increase background faster than target signal.
  4. Clarify again if samples become viscous or particulate.

3Run SDS-PAGE

  1. Choose gel percentage by target size.
  2. Avoid distorted lanes, since weak targets become harder to interpret in smeared samples.
  3. Run the target region with enough separation from abundant nearby proteins.
  4. Do not compensate for low abundance by overloading until enrichment has been considered.

4Transfer and check before probing

  1. Use transfer settings based on target size.
  2. Stain the membrane after transfer and document the target molecular weight region.
  3. If the positive control is weak, troubleshoot transfer before antibody dilution.
  4. For small or large low-abundance targets, use the matching small- or large-protein transfer approach.

5Increase detection sensitivity in order

  1. Use overnight primary incubation at 4°C.
  2. Use a sensitive ECL or fluorescent detection reagent.
  3. Increase Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody concentration only after confirming transfer and positive control.
  4. Avoid increasing secondary antibody first if background is already present.

Controls to include

  • Strong known positive control to confirm the assay can detect the target.
  • No-primary or secondary-only control if background is a concern.
  • Fraction marker or enrichment control if enrichment is used.
  • Loading control or total protein stain suited to the fraction or sample type.

Small protein, histone, or peptide Western blot protocol

For histones, peptides, small cytokines, and proteins below about 20 kDa.

  • Use this workflow when: The target is a small protein, histone, peptide-sized target, or low molecular weight modification-sensitive protein.
  • Optimize first: High-percentage gel selection, membrane pore size, transfer time, methanol level, and prevention of over-transfer.

Before you start

  • Use a high-percentage gel or Tris-Tricine gel for small target resolution.
  • Prepare 0.2/0.22 um membrane to reduce blow-through.
  • Use lower sample loads for abundant histones. For histone extraction from mammalian cells or tissues, consider EpiQuik Total Histone Extraction Kit (OP-0006).
  • Plan total H3, total H4, total histone, peptide, or recombinant control when appropriate.

Protocol

1Prepare the small-protein sample

  1. Use histone extract, enriched fraction, purified peptide, or total lysate depending on the target.
  2. For histone extracts, start around 0.5-5 ug per lane.
  3. For total lysate small targets, use the lowest load that still gives visible target signal.
  4. Avoid excessive loading because small abundant proteins can saturate quickly.

2Denature

  1. Add sample buffer to 1X final concentration.
  2. Heat at 95°C for 5 minutes unless the antibody or sample type requires another condition.
  3. Briefly centrifuge before loading.
  4. Keep peptide or histone controls prepared consistently with samples.

3Run a small-protein gel

  1. Use 15-18% gel or Tris-Tricine gel for very small targets.
  2. Run until the small target is separated from the dye front and nearby bands.
  3. Use a ladder that covers low molecular weight markers.
  4. If small bands compress, change gel chemistry before changing antibody conditions.

4Transfer gently

  1. Use 0.2/0.22 um PVDF or nitrocellulose.
  2. Use shorter or gentler transfer than routine proteins.
  3. Avoid SDS in transfer buffer unless specifically validated.
  4. If signal is missing, check for membrane blow-through before increasing antibody.

5Block, probe, and normalize

  1. Use blocker compatible with the antibody. For many histone PTM antibodies, start with BSA in TBST.
  2. Probe with target or modification-specific antibody.
  3. Normalize histone targets to total H3/H4, total histone, or total protein signal when appropriate.
  4. Use short exposures first because histone signals can saturate.

Controls to include

  • Total H3, total H4, or total histone control for histone targets.
  • Modified peptide, recombinant histone, or known positive sample for PTM targets.
  • Unmodified peptide or untreated sample when testing modification specificity.
  • Transfer retention check because small proteins can pass through membranes.

Large protein Western blot protocol

For proteins above about 150 kDa, high molecular weight targets, or proteins that transfer poorly.

  • Use this workflow when: The target is a large protein that transfers slowly or resolves poorly on a standard gel.
  • Optimize first: Gel percentage, transfer time, membrane choice, sample denaturation, and reducing conditions.

Before you start

  • Use a low-percentage or gradient gel suitable for high molecular weight separation.
  • Plan wet transfer with cooling and longer transfer time.
  • Prepare reduced-methanol transfer buffer when needed.
  • Plan membrane stain and post-transfer gel stain to check whether protein remains in the gel.

Protocol

1Prepare clean lysate

  1. Lyse on ice and avoid harsh handling that promotes aggregation.
  2. Clarify thoroughly before loading.
  3. Start with 10-30 ug total protein per lane depending on target abundance.
  4. Avoid viscous or particulate samples, which can trap large proteins near the well.

2Denature with care

  1. Use reducing sample buffer unless non-reducing conditions are required.
  2. Start with 95°C for 5 minutes for routine large soluble proteins.
  3. If the protein aggregates or stays near the well, test 37-70°C for 10-30 minutes.
  4. Briefly centrifuge before loading.

3Run a large-protein gel

  1. Use 4-8% gel or 4-15% gradient gel depending on target size.
  2. Run longer at appropriate voltage to improve high molecular weight separation.
  3. Use a high molecular weight ladder.
  4. If the target remains compressed near the top, lower the gel percentage before changing antibody conditions.

4Use extended wet transfer

  1. Use wet transfer with cooling instead of a short routine transfer.
  2. Start with 30 V overnight or another validated extended wet-transfer condition.
  3. Reduce methanol to 5-10% if large protein transfer is poor.
  4. Consider 0.025-0.05% SDS only after optimizing methanol and transfer time.

5Probe and check transfer first

  1. Block with milk or datasheet-recommended blocker.
  2. Use antibody conditions from the datasheet when available.
  3. If signal is weak, stain the post-transfer gel to determine whether target remained in the gel.
  4. Optimize transfer before increasing antibody concentration.

Controls to include

  • High molecular weight ladder.
  • Known positive lysate for the large target.
  • Ponceau S or total protein stain on membrane.
  • Post-transfer gel stain when transfer efficiency is uncertain.

Membrane or hydrophobic protein Western blot protocol

For membrane-enriched, hydrophobic, multi-pass, organelle, or detergent-sensitive proteins.

  • Use this workflow when: The target is hydrophobic, multi-pass, poorly soluble, or found mainly in membrane fractions.
  • Optimize first: Lysis buffer detergents, sample heating conditions, membrane fraction enrichment, gel percentage, and transfer efficiency.

Before you start

  • Choose lysis or extraction conditions strong enough to solubilize the target.
  • Prepare membrane fraction and cytosolic marker controls when fractionation is used.
  • Plan a denaturation comparison if boiling may cause aggregation.
  • Confirm detergent compatibility with SDS-PAGE and antibody detection.

Protocol

1Extract the membrane target

  1. Use detergent conditions appropriate for the membrane protein and sample type.
  2. Keep samples cold and include fresh inhibitors.
  3. Clarify insoluble debris after extraction.
  4. If fractionating, keep membrane and cytosolic fractions clearly separated.

2Normalize and test denaturation

  1. Normalize by protein concentration or fraction-equivalent loading.
  2. Start with standard reducing sample buffer.
  3. If boiling causes aggregation, test 37-70°C for 10-30 minutes.
  4. Avoid repeated freeze-thaw cycles, which can worsen aggregation.

3Run SDS-PAGE

  1. Choose gel percentage by target size.
  2. Watch for material remaining in the well or broad smearing.
  3. If migration is poor, optimize extraction and denaturation before changing antibody conditions.
  4. Use fraction marker lanes when possible.

4Transfer

  1. Use transfer settings based on target size.
  2. Use 0.45 um membrane for most membrane proteins above about 20 kDa.
  3. Check transfer before blocking.
  4. Optimize transfer only after sample solubilization is acceptable.

5Probe and reduce background

  1. Start with datasheet-recommended blocker or 5% milk in TBST for routine targets.
  2. Increase wash stringency if hydrophobic samples create broad background.
  3. Confirm signal is enriched in the membrane fraction when fractionation is used.
  4. If target is weak, troubleshoot extraction before increasing secondary antibody.

Controls to include

  • Membrane fraction marker to confirm enrichment.
  • Cytosolic marker to check fraction purity.
  • Positive membrane-rich sample.
  • No-primary control if hydrophobic background is high.

Phospho-specific or labile PTM Western blot protocol

For phosphorylation or other modification-specific targets that require careful sample handling and controls.

  • Use this workflow when: You are detecting phosphorylation, acetylation, methylation, ubiquitination, or another modification-specific target.
  • Optimize first: Inhibitor protection, treatment controls, blocking buffer, antibody specificity controls, and non-saturated exposure.

Before you start

  • Add phosphatase and protease inhibitors fresh to lysis buffer.
  • Keep samples cold and process quickly.
  • Use BSA in TBST as the starting blocker unless the antibody datasheet recommends otherwise.
  • Prepare total target, stimulated/unstimulated, inhibitor-treated, or phosphatase-treated controls when possible.

Protocol

1Lyse quickly with inhibitors

  1. Chill samples, tubes, and buffer before lysis.
  2. Add inhibitors immediately before use.
  3. Lyse and clarify samples at 4°C.
  4. Avoid delays, repeated freeze-thaw cycles, and prolonged handling.

2Normalize and denature

  1. Quantify protein and normalize all samples.
  2. Add sample buffer promptly after lysis.
  3. Heat at 95°C for 5 minutes unless the antibody datasheet recommends another condition.
  4. Keep the handling time consistent across treatment groups.

3Run SDS-PAGE

  1. Choose gel percentage by the total target protein size.
  2. Use better-resolving conditions if a modification-dependent mobility shift is expected.
  3. Run treated and untreated samples together.
  4. Avoid overloading, which can obscure subtle PTM-dependent changes.

4Transfer

  1. Transfer based on target size and confirm transfer before probing.
  2. Use membrane stain to verify consistent lane transfer.
  3. If phospho signal is weak, confirm total target before increasing phospho antibody.
  4. Use small- or large-protein transfer settings if target size requires them.

5Block, probe, and confirm specificity

  1. Block and dilute antibody in 3-5% BSA in TBST unless datasheet recommends otherwise.
  2. Probe phospho/PTM signal and total target protein.
  3. Confirm signal changes with stimulation, inhibition, or phosphatase treatment when possible.
  4. Switch blocker before assuming the antibody failed if background is high.

Controls to include

  • Total target protein to confirm expression and loading.
  • Stimulated and unstimulated samples to show expected modification change.
  • Phosphatase-treated sample when applicable.
  • Pathway inhibitor or treatment control when applicable.

IP, co-IP, or pulldown Western blot protocol

For IP-Western analysis of immunoprecipitated, co-IP, pulldown, bead-bound, or enriched eluate samples. This Western blot workflow assumes the IP or pulldown capture step has already been performed.

  • Use this workflow when: The immunoprecipitation, co-IP, pulldown, or enrichment step has already been performed and the eluate or bead-bound fraction will be analyzed by Western blot.
  • Optimize first: Input and IgG controls, heavy-chain and light-chain interference, elution method, sample loading, and detection antibody strategy.

Before you start

  • Plan lane order before starting: input, flow-through, wash, eluate, IgG IP, and beads-only when possible.
  • Check whether the target overlaps IgG heavy chain near 50 kDa or light chain near 25 kDa.
  • Choose a detection strategy that avoids IP antibody chain interference when needed.
  • Keep input and control samples for interpretation.

Protocol

1Collect IP fractions

  1. Save input before IP.
  2. Save flow-through and wash fractions if capture efficiency matters.
  3. Elute target from beads under conditions compatible with downstream Western blotting.
  4. Include IgG IP and beads-only control when possible.

2Prepare eluate for loading

  1. Add sample buffer to eluate or bead-bound sample according to the IP method.
  2. Boil beads in sample buffer only if compatible with the experiment.
  3. Briefly centrifuge and load the supernatant without bead carryover.
  4. Load input as a small percentage of starting material and eluate as an equivalent fraction.

3Run SDS-PAGE

  1. Choose gel percentage by target size.
  2. Account for IgG heavy chain near 50 kDa and light chain near 25 kDa.
  3. If the target overlaps IgG chains, use detection designed to avoid chain interference.
  4. Separate input, control IP, and test IP lanes clearly.

4Transfer

  1. Use transfer settings based on target size.
  2. Confirm transfer before blocking.
  3. If sample amount is limited, avoid unnecessary membrane stripping.
  4. Use small- or large-target transfer changes only when target size requires them.

5Probe and interpret enrichment

  1. Use light-chain-specific, conformation-specific, or directly conjugated detection if IgG chain interference is expected.
  2. Compare test IP to IgG IP and beads-only control.
  3. Use input to confirm that the target was present before IP.
  4. Troubleshoot enrichment before assuming Western blot antibody failure.

Controls to include

  • Input to show the target was present before IP.
  • Flow-through and wash to evaluate capture and wash loss.
  • IgG IP to measure non-specific antibody pulldown.
  • Beads-only IP to measure non-specific bead binding.

Secreted protein or conditioned media Western blot protocol

For conditioned media, culture supernatant, serum-containing media, or concentrated secreted protein samples.

  • Use this workflow when: The target is secreted into conditioned media, serum-free media, plasma, serum, or another low-protein extracellular sample.
  • Optimize first: Sample concentration, matrix background, loading normalization, positive controls, and transfer/detection sensitivity.

Before you start

  • Clarify media to remove cells and debris before concentration or loading.
  • Choose a normalization basis: volume, cell number, viability, total protein, or another experiment-specific factor.
  • Concentrate media if the target is expected to be dilute.
  • Include blank media and conditioned-media positive control when possible.

Protocol

1Clarify and normalize media

  1. Centrifuge conditioned media to remove cells and debris.
  2. Concentrate the media if target abundance is low.
  3. Normalize by volume, cell number, viability, total protein, or collection time.
  4. Record the normalization basis so lanes can be compared.

2Prepare sample buffer

  1. Add sample buffer after concentration or cleanup.
  2. Use reducing or non-reducing conditions based on the target and antibody.
  3. If concentrated media contains high salt, desalt or clean up before loading.
  4. Avoid overloading serum proteins if serum-containing media is used.

3Run SDS-PAGE

  1. Choose gel percentage by target size.
  2. Run blank media, conditioned media, and positive control on the same gel.
  3. If serum proteins dominate the lane, reduce load or use serum-free collection when feasible.
  4. If migration is distorted, check salt and concentration method.

4Transfer

  1. Transfer based on target size.
  2. Use 0.45 um membrane for most routine secreted proteins above about 20 kDa.
  3. Confirm transfer with membrane stain.
  4. If target is small, use small-protein transfer settings.

5Probe and compare to media controls

  1. Block with datasheet-recommended blocker or 5% milk for routine targets.
  2. Increase washing if serum or media components create background.
  3. Compare conditioned media against blank media control.
  4. Use viability data to distinguish secretion from protein release due to cell death.

Controls to include

  • Blank media to identify media-derived background.
  • Conditioned-media positive control or purified target.
  • Cell viability control to help distinguish secretion from cell lysis.
  • Normalization record for volume, cell number, viability, or protein basis.
Open full WB protocol

Choose Your ELISA Format

Use this ELISA protocol guide to choose and run a plate-based immunoassay workflow. Select sandwich, indirect, direct, or competitive ELISA to view detailed benchtop starting conditions, including coating, blocking, washing, antibody incubation, detection, plate reading, controls, and troubleshooting.

Sandwich ELISA protocol

Use this workflow when a target is captured by one antibody and detected by a second antibody that recognizes a different epitope.

  • Use this format when: Quantifying low-abundance soluble antigens in complex biological matrices where a compatible matched antibody pair is available.
  • Optimize first: Capture antibody concentration, detection antibody concentration, sample dilution, blocking buffer, wash stringency, standard curve range, and matrix effects.
Sample types
  • Best sample types: Serum, plasma, saliva, culture supernatants, and tissue homogenates.
  • Best for: Quantifying low-abundance antigens, such as cytokines, hormones, and biomarkers, in complex biological matrices.
  • Accuracy note: Sandwich ELISA is usually not the best choice for very small or single-epitope analytes because two non-overlapping antibody binding sites are needed.
1
Prepare plate map and reagents
Plan standards, blanks, negative controls, positive controls, and samples before starting. Bring coated plate, standards, samples, wash buffer, blocking buffer, detection antibody, enzyme conjugate, substrate, and stop solution to room temperature unless the kit manual says otherwise. For manual coating, use a high-binding 96-well ELISA plate.
2
Coat capture antibody
Dilute capture antibody in carbonate-bicarbonate coating buffer, pH 9.6, or the buffer specified for the antibody pair. Add 100 uL per well. Cover the plate and incubate overnight at 4 degrees C, or 2 hours at room temperature as a faster starting condition. Aspirate the coating solution after incubation.
3
Wash and block
Wash 3 times with 250-300 uL per well 1x PBST or TBST. Add 200 uL per well blocking buffer, such as 1-5 percent BSA, casein, or nonfat dry milk in PBS/TBS. Incubate 1 hour at room temperature. Wash 3 times.
4
Add standards and samples
Prepare standards in duplicate or triplicate using the recommended diluent. Dilute samples so readings fall within the linear range. For cell lysates or tissue homogenates, prepare clarified samples and include fresh protease inhibitor during upstream extraction when compatible with the assay, such as R-1101 Protease Inhibitor Cocktail. Add 100 uL per well and incubate 1-2 hours at room temperature, or overnight at 4 degrees C for low-abundance targets. Wash 4-5 times.
5
Add detection antibody
Dilute detection antibody in assay diluent. If this product is used as the detection antibody, add Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody at the datasheet-recommended dilution or start at 0.25-2 ug/mL for optimization. Add 100 uL per well and incubate 1 hour at room temperature. Wash 4-5 times.
6
Add enzyme conjugate and substrate
Add 100 uL per well HRP-conjugated secondary antibody, or R-1098 Streptavidin: HRP Conjugate when the detection antibody or binding reagent is biotin-labeled. Incubate 30-60 minutes at room temperature protected from light when needed. Wash 5 times, then add 100 uL TMB substrate per well.
7
Stop and read
Develop color for 5-20 minutes until the standard curve is visible but not saturated. Add 50-100 uL stop solution per well and read absorbance at 450 nm, with 570 or 620 nm correction if available. Analyze standards with a 4-parameter logistic curve when appropriate.
ControlsInclude blank wells, zero standard, standard curve, matrix control, no-sample control, and positive sample when available.
Wash consistencyMost high background in sandwich ELISA comes from insufficient washing, over-concentrated detection antibody, or incompatible blocking buffer.

Direct ELISA protocol

Use this workflow when the target is immobilized or captured and detected directly by a labeled antibody, direct detection reagent, or kit-specific detection system.

  • Use this format when: A direct detection kit, directly labeled antibody, purified antigen, semi-purified antigen, or target-specific direct detection reagent is available.
  • Optimize first: Coating or sample input, target purity, detection reagent amount, incubation time, wash stringency, and substrate development time.
Sample types
  • Best sample types: Purified or semi-purified antigens, recombinant proteins, and clarified cell or tissue lysates when target coating and background are acceptable.
  • Best for: Screening antibody specificity or confirming the presence of an antigen in an isolated, high-purity state.
  • Accuracy note: Direct ELISA is simple and fast, but it is less ideal for crude complex matrices because non-target proteins can compete for plate binding and increase background.
1
Prepare plate and samples
Plan standards, blanks, negative controls, positive controls, and sample dilutions. For manual direct ELISA, use purified or semi-purified antigen, recombinant protein, or clarified lysate only when plate coating and background are acceptable. Add 100 uL per well of standards or samples according to the kit format or coating requirement.
2
Immobilize target if required
For coated-plate direct ELISA, incubate antigen or sample in the well overnight at 4 degrees C or 2 hours at room temperature. For direct target-detection kits, follow the kit manual for binding, capture, or sample incubation conditions.
3
Wash and block
Wash 3 times with 250-300 uL per well wash buffer. Add 200 uL per well blocking buffer for 30-60 minutes if the assay format requires blocking. Some direct detection kits include a proprietary blocking or assay buffer that should be used instead.
4
Add direct detection reagent
Add 100 uL per well of the directly labeled antibody, direct detection reagent, or enzyme-conjugated target-specific reagent. If using a directly conjugated antibody, start with the datasheet-recommended dilution or test 0.1-1 ug/mL. Incubate 30-90 minutes at room temperature protected from light when appropriate.
5
Wash and develop
Wash 4-5 times to reduce background. Add 100 uL per well substrate. Develop until standards or positive controls show clear separation without saturation.
6
Stop and read
Add 50-100 uL stop solution when using TMB and read at 450 nm. Use the correction wavelength recommended for the plate reader if available. Compare results to the standard curve or kit-specific calculation method.
Direct detection kitsSome EpigenTek assay kits use direct target-detection formats for small molecules, DNA/RNA damage markers, or epigenetic modifications. In those cases, the kit manual overrides generic antibody-only workflow assumptions.
Background controlInclude blank, no-target, no-detection-reagent, and positive control wells when compatible with the kit or assay design.

Indirect ELISA protocol

Use this workflow to detect antibody binding to an immobilized antigen or target molecule using an unconjugated primary antibody and labeled secondary antibody.

  • Use this format when: Measuring antibody binding, total antibody concentration, titer, or immune response to a coated antigen.
  • Optimize first: Antigen coating concentration, sample or antibody dilution, blocking buffer, secondary antibody dilution, and wash stringency.
Sample types
  • Best sample types: Serum, plasma, saliva, and other body fluids.
  • Best for: Measuring total antibody concentrations, titers, or screening immune responses against specific pathogens or antigens.
  • Accuracy note: Match the secondary antibody to the sample antibody species and isotype, and validate saliva or other non-serum matrices before relying on quantitative comparisons.
1
Coat antigen or target
Dilute purified antigen, peptide, protein, histone, modified target, cell lysate, or other coating material in carbonate-bicarbonate buffer or PBS. Add 100 uL per well. A common starting range is 0.5-5 ug/mL for purified protein or peptide. If the antigen or capture reagent is biotinylated, a streptavidin-coated plate such as R-1102 Streptavidin-Coated Strip Microwell Plate may be used instead of passive coating. Incubate overnight at 4 degrees C or 2 hours at room temperature.
2
Wash and block
Aspirate coating solution and wash 3 times with 250-300 uL per well PBST or TBST. Add 200 uL per well blocking buffer. Incubate 1 hour at room temperature, then wash 3 times.
3
Add primary antibody
Dilute Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody in assay diluent. Start with the datasheet-recommended dilution when available; otherwise test a dilution series such as 1:250, 1:500, 1:1000, and 1:2000. Add 100 uL per well and incubate 1 hour at room temperature or overnight at 4 degrees C for weak targets.
4
Wash after primary antibody
Wash 4-5 times with 250-300 uL per well wash buffer. Tap the inverted plate on clean absorbent material after the final wash to remove residual liquid without drying the wells.
5
Add enzyme-conjugated secondary antibody
Dilute HRP- or AP-conjugated secondary antibody in assay diluent, commonly 1:2000-1:10000 depending on supplier and signal strength. For common mouse or rabbit primary antibodies, host-matched options include A12003 HRP-Goat Anti-Mouse Secondary Antibody or A12004 HRP-Goat Anti-Rabbit Secondary Antibody. Add 100 uL per well and incubate 30-60 minutes at room temperature.
6
Develop signal
Wash 5 times. Add 100 uL substrate per well. For HRP/TMB, develop 5-20 minutes protected from strong light, then stop with 50-100 uL stop solution and read at 450 nm.
7
Interpret results
Subtract blank background and compare signal across antigen-coated, uncoated, no-primary, and no-secondary controls. High signal in no-primary wells suggests secondary antibody or blocking-related background.
Antigen coatingToo much coated antigen can increase background. If the signal is high but nonspecific, reduce coating concentration before changing everything else.
Primary antibody variableThe product page can replace the primary antibody variable in this protocol with the selected antibody name.

Competitive ELISA protocol

Use this workflow when sample target competes with immobilized or labeled target for antibody or binding reagent.

  • Use this format when: The analyte is small, has one dominant epitope, or is measured by competition rather than two-antibody sandwich capture.
  • Optimize first: Sample dilution, competitor concentration, antibody concentration, incubation time, standard curve range, and matrix interference.
Sample types
  • Best sample types: Serum, plasma, saliva, culture supernatants, and crude biological fluids.
  • Best for: Detecting small molecules, hormones, or targets with only a single antigenic epitope.
  • Accuracy note: Competitive ELISA produces an inverse signal, so higher analyte levels usually give lower absorbance or fluorescence.
1
Prepare standards and samples
Prepare a full standard curve using the assay diluent recommended for the target. For serum, plasma, saliva, culture supernatants, or crude biological fluids, dilute samples into the same matrix when possible. Competitive assays often need careful sample dilution because high target concentration produces lower signal.
2
Add standards, samples, and competitor
Add 50-100 uL per well of standards or samples according to the assay design. Add competitor, tracer, or labeled target reagent if the protocol requires a pre-mix. Mix gently without splashing.
3
Add antibody or binding reagent
Add the antibody or target-specific binding reagent at the optimized concentration. When using an antibody-based format, add Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody only if the product is the intended competitive binding antibody and follow the datasheet dilution first.
4
Incubate competition reaction
Incubate 1-2 hours at room temperature, or as specified by the kit manual. Keep incubation times consistent across the plate because competitive assays are especially sensitive to timing differences.
5
Wash thoroughly
Wash 4-5 times with 250-300 uL per well wash buffer. Avoid drying the wells. Inconsistent washing can distort the inverse standard curve.
6
Develop and read
Add 100 uL substrate per well and develop until the low-analyte standards produce strong but unsaturated signal. Stop and read at the appropriate wavelength. Fit the data using the curve model recommended by the kit or assay format.
Signal directionIn competitive ELISA, stronger sample target usually means lower signal. Label graphs and reports clearly to avoid reversed interpretation.
TimingUse a multichannel pipette and consistent timing between rows when possible.
Open full ELISA protocol

Chromatin Immunoprecipitation (ChIP) Protocol

Use this guided ChIP protocol page to choose a cross-linking ChIP, native ChIP, ChIP-qPCR, or ChIP-seq preparation workflow. The steps below are practical starting conditions for antibody-based enrichment of chromatin-associated DNA. Optimize fixation, fragmentation, antibody amount, bead type, wash stringency, and downstream detection for each target and sample type.

For kit-supported workflows, choose a ChIP enrichment kit, qPCR reagent, or ChIP-seq library preparation kit that matches your starting material, input amount, and downstream readout.

Typical starting inputs

  • Histone marks: 1 x 105 to 1 x 106 cells per IP may be sufficient after optimization.
  • Transcription factors: start with 1 x 106 to 1 x 107 cells per IP.
  • Low-abundance cofactors: use higher input, stronger controls, and replicate IPs.

Native ChIP protocol

Use native ChIP for histones and many histone modifications when preserving native nucleosome structure is preferred and formaldehyde crosslinking is not required.

  • Use this format when: The target is a histone protein or histone modification with strong chromatin association.
  • Optimize first: MNase digestion, nucleosome fragment range, antibody amount, and salt conditions during immunoprecipitation.
1
Collect cells without crosslinking
Start with 1 x 106 to 5 x 106 cells per IP for histone targets as a practical starting range. Wash cells twice with cold PBS and keep all steps cold unless digestion conditions require warmth.
2
Prepare nuclei
Resuspend the cell pellet in hypotonic or nuclear isolation buffer containing protease inhibitors. Incubate on ice 10 minutes. Dounce gently if needed. Pellet nuclei at 800-1,000 x g for 5 minutes at 4°C.
3
Digest chromatin with MNase
Resuspend nuclei in MNase digestion buffer containing calcium. Add a titrated amount of micrococcal nuclease and incubate at 37°C for 5-20 minutes. Start with a small pilot digestion to identify conditions that produce mostly mono- and oligonucleosomes.
4
Stop digestion
Add EDTA to 5-10 mM final concentration to stop MNase activity. Place samples on ice immediately. Avoid prolonged digestion after the stop step.
5
Release soluble chromatin
Lyse nuclei gently with native ChIP lysis or IP buffer containing 0.1-0.5 percent NP-40 or Triton X-100 and physiological salt. Incubate on ice 10 minutes, then clarify at 12,000 x g for 10 minutes at 4°C.
6
Check nucleosome size and save input
Remove 20-50 uL chromatin for DNA purification and size check. A useful native ChIP digestion often includes mono-, di-, and tri-nucleosome DNA with a major range around 150-600 bp. Save 1-10 percent input.
7
Pre-clear chromatin
Incubate chromatin with 20-40 uL blocked protein A/G beads for 30-60 minutes at 4°C with rotation. Transfer the pre-cleared supernatant to a new tube.
8
Add histone or PTM antibody
Add 1-5 ug Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody per IP as a starting range. For abundant histone marks, lower antibody and lower chromatin input may be sufficient. Rotate 2-4 hours or overnight at 4°C.
9
Capture with beads
Add 30-50 uL washed, blocked protein A/G beads and rotate 1-2 hours at 4°C. Use magnetic separation or low-speed spin collection depending on bead type.
10
Wash under native conditions
Wash 4-6 times with cold native ChIP wash buffer. Start with 1 mL per wash and 3-5 minutes rotation. Increase salt or detergent only if background is high and recovery remains acceptable.
11
Elute DNA-protein complexes
Elute with 100-200 uL elution buffer compatible with DNA recovery, commonly SDS-containing elution buffer or proteinase K digestion buffer. Treat with proteinase K at 55-65°C for 1-2 hours.
12
Purify DNA and quantify enrichment
Purify DNA and elute in 30-50 uL. Analyze by qPCR using input-normalized percent input or fold enrichment over IgG. For sequencing, confirm DNA yield and library compatibility before pooling samples.
Best fitNative ChIP is usually strongest for histone marks and may not preserve weak or transient transcription factor binding.
MNase titrationMNase digestion is sample dependent. Always optimize enzyme amount and digestion time before large experiments.

ChIP-qPCR analysis protocol

Use ChIP-qPCR to validate enrichment at defined genomic loci after native or cross-linking ChIP.

  • Use this format when: Testing a known locus, validating ChIP enrichment, optimizing a new ChIP antibody, or confirming ChIP-seq findings.
  • Relevant ChIP-qPCR kits: Pair ChIP-enriched DNA with the EpiQuik Quantitative PCR Fast Kit for fast quantitative real-time PCR analysis of ChIP DNA. Use a ChIP enrichment kit such as EpiQuik ChIP Kit or ChromaFlash High-Sensitivity ChIP Kit upstream.
  • Optimize first: Primer specificity, amplicon size, qPCR efficiency, positive and negative loci, input dilution, and replicate consistency.
1
Design ChIP-qPCR primers
Design primers for 70-150 bp amplicons centered on the expected binding or modification region. Include at least one positive locus and one negative locus when possible.
2
Validate primer efficiency
Test primers on input DNA dilution series. Aim for a single melt peak and similar amplification efficiency across primer sets. Redesign primers if primer dimers or non-specific products appear.
3
Dilute ChIP and input DNA
Use 1-2 uL purified ChIP DNA per 10-20 uL qPCR reaction as a starting point. Dilute input DNA so Ct values fall in the linear range and are not several cycles earlier than the ChIP samples.
4
Set up qPCR reactions
For each 20 uL reaction, combine 10 uL 2x qPCR master mix, primer mix at validated concentration, 1-2 uL DNA, and nuclease-free water. The EpiQuik Quantitative PCR Fast Kit can be used for ChIP DNA qPCR analysis. Run technical duplicates or triplicates for input, target IP, IgG IP, and controls.
5
Use consistent controls
Include no-template controls, input DNA, IgG IP DNA, positive control IP DNA when available, and negative genomic loci. If using Symmetric DiMethyl-Histone H3-R8 Polyclonal Antibody for a new target, include a known responsive biological condition when possible.
6
Calculate percent input
Correct input Ct for dilution, then calculate percent input for each IP. A common formula is 100 x 2^(adjusted input Ct - IP Ct). Apply the same formula to target IP and IgG IP.
7
Calculate fold enrichment
Calculate enrichment over IgG or over a negative locus when appropriate. Report the normalization method clearly because percent input and fold enrichment answer different questions.
8
Interpret enrichment with controls
A useful ChIP-qPCR result should show enrichment at expected loci, low IgG background, acceptable input amplification, and reproducibility across biological replicates. Weak signal at all loci usually points to chromatin, antibody, or wash stringency issues.
Amplicon sizeShort amplicons perform better with fragmented ChIP DNA. Long amplicons can underperform after heavy shearing.
Negative lociNegative loci help distinguish real locus-specific enrichment from antibody or bead background.

ChIP-seq preparation and QC protocol

Use this workflow after native or cross-linking ChIP when the enriched DNA will be used for sequencing library preparation.

  • Use this format when: ChIP DNA will be converted into sequencing libraries for genome-wide mapping.
  • Relevant ChIP-seq kit: For an all-in-one ChIP and ChIP-seq library preparation workflow, see the EpiNext ChIP-Seq High-Sensitivity Kit (Illumina).
  • Optimize first: Fragment size, DNA recovery, adapter dilution, PCR cycle number, duplicate rate, and positive or negative qPCR QC before sequencing.
1
Confirm ChIP enrichment before library prep
Run ChIP-qPCR on at least one positive and one negative locus before committing samples to sequencing. Proceed when target IP shows clear enrichment over IgG and negative loci.
2
Quantify ChIP DNA carefully
Use a sensitive fluorometric method or library prep kit guidance for low-input DNA. ChIP DNA may be below the accurate range of standard absorbance methods.
3
Assess fragment size
Check purified ChIP DNA or library intermediates by Bioanalyzer, TapeStation, Fragment Analyzer, or gel-based method. Many ChIP-seq workflows use enriched DNA around 150-500 bp before library amplification.
4
Prepare library with low-input conditions
Use a ChIP-seq compatible low-input DNA library kit such as the EpiNext ChIP-Seq High-Sensitivity Kit (Illumina). Follow kit-specific end repair, A-tailing, adapter ligation, cleanup, and PCR conditions. Dilute adapters when DNA input is low to reduce adapter dimers.
5
Optimize PCR cycle number
Use the fewest PCR cycles that produce enough library for sequencing. Excess PCR can increase duplicate reads, skew enrichment, and create library artifacts.
6
Clean up and size select library
Use magnetic bead cleanup or kit-recommended size selection to remove primers, enzymes, salts, and adapter dimers. Verify final library size distribution before pooling.
7
Quantify final library
Quantify libraries using qPCR-based library quantification or another sequencing-compatible method. Normalize samples before pooling.
8
Review sequencing design
Include biological replicates, matching input controls, and IgG controls when informative. For broad histone marks, plan sufficient depth and appropriate peak-calling settings. For narrow transcription factor peaks, prioritize enrichment and replicate consistency.
Library artifactsAdapter dimer and high PCR duplication often indicate too little input DNA, excess adapter, or too many PCR cycles.
ReplicatesBiological replicate consistency is more useful than a single high-depth library with poor ChIP enrichment.
Open full ChIP protocol

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