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Base Cat #A73465

ATOX1 Polyclonal AntibodyKO Validated

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Covers ATOX1 as a marker or regulator in membrane transport, trafficking, and cellular localization.

We recommend using an alternative (Catalog #A81610): ATOX1 Polyclonal Antibody
Application: IFWB
Clonality: Polyclonal
Conjugate: Unconjugated
Host: Rabbit
Isotype: IgG
Knockout Validated: Yes
Purification: Affinity Purified
Reactivity: Human
100% Guarantee: 6 months
Application Guarantee for AntibodiesRecombinant Rabbit Monoclonal Abs
Catalog No.SizePriceQty
A73465-100100 µl $368.00 
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Availability: Discontinued 

Product Overview

Background
This gene encodes a copper chaperone that plays a role in copper homeostasis by binding and transporting cytosolic copper to ATPase proteins in the trans-Golgi network for later incorporation to the ceruloplasmin. This protein also functions as an antioxidant against superoxide and hydrogen peroxide, and therefore, may play a significant role in cancer carcinogenesis. Because of its cytogenetic location, this gene represents a candidate gene for 5q-syndrome.

Description
ATOX1 Polyclonal Antibody. Unconjugated. Raised in: Rabbit.

Formulation
Buffer: PBS with 0.02% sodium azide, 50% glycerol, pH7.3.

Specificity
Human

Isotype
IgG

Uniprot ID
O00244

Purification
Affinity Purified

Immunogen
Recombinant protein of human ATOX1.

Storage
Shipped at 4°C. Store at -20°C. Avoid freeze / thaw cycles.

Alternative Names
ATOX1; ATX1; HAH1

Application
WB, IF; Recommended dilution: WB 1:500 - 1:2000, IF 1:50 - 1:100

Datasheet & SDS

PDF
Datasheet/User Guide: A73465 If the above file is not available online, you can request a copy by contacting us.

Applications

Suggested protocols for ATOX1 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 ATOX1 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, ATOX1 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 ATOX1 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 ATOX1 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 IF/ICC Workflow

Use this immunofluorescence (IF) and immunocytochemistry (ICC) protocol guide to run fluorescent antibody staining in cultured cells, tissue sections, multiplex panels, and phospho/PTM target workflows.

Cultured cell IF/ICC protocol

Use this workflow for fluorescent antibody staining of adherent or suspension cells on coverslips, chamber slides, or imaging plates.

  • Use this format when: Detecting protein localization, nuclear markers, cytoplasmic proteins, histone marks, signaling proteins, or cell morphology in cultured cells.
  • Optimize first: Fixation method, permeabilization strength, blocking buffer, primary antibody dilution, and imaging exposure.
1
Plate cells
Seed cells onto sterile coverslips, chamber slides, or imaging plates so they reach 50-80 percent confluence on staining day. For coverslips in a 24-well plate, use enough medium to fully cover the cells.
2
Fix cells
Remove medium and gently rinse once with PBS. Add 4 percent paraformaldehyde in PBS for 10-15 minutes at room temperature. For some cytoskeletal, membrane, or epitope-sensitive targets, compare with cold methanol fixation for 5-10 minutes at -20 degrees C.
3
Wash and permeabilize
Wash 3 times with PBS, 5 minutes each. Permeabilize with 0.1-0.3 percent Triton X-100 in PBS for 5-10 minutes at room temperature. For nuclear histone or chromatin targets, 0.2-0.5 percent Triton X-100 can be tested. For membrane targets, reduce or omit detergent.
4
Block nonspecific binding
Add blocking buffer, such as 1-5 percent BSA or 5 percent normal serum in PBS with 0.05-0.1 percent Tween-20. Incubate 30-60 minutes at room temperature. Use serum from the host species of the secondary antibody when practical.
5
Add primary antibody
Dilute ATOX1 Polyclonal Antibody in blocking buffer. Start with the datasheet-recommended IF/ICC dilution or test 1:100, 1:250, and 1:500. Add enough volume to cover the sample, such as 100-200 uL per coverslip or 50-100 uL per chamber well. Incubate 1 hour at room temperature or overnight at 4 degrees C in a humidified chamber.
6
Wash after primary antibody
Wash 3 times with PBS or PBST, 5 minutes each, using gentle rocking. Do not allow cells to dry.
7
Add fluorescent secondary antibody
Dilute fluorophore-conjugated secondary antibody in blocking buffer, commonly 1:500-1:1000. Add enough volume to cover the cells and incubate 45-60 minutes at room temperature protected from light.
8
Counterstain and mount
Wash 3 times. Add DAPI or another nuclear counterstain for 5 minutes if desired, then wash once. Mount coverslips with antifade mounting medium and cure according to the mountant instructions.
9
Image and document settings
Acquire images using the same exposure, laser power, gain, and offset for samples that will be compared. Include no-primary and secondary-only controls to evaluate background.
Fixation choiceParaformaldehyde preserves morphology well. Methanol can improve some cytoskeletal or nuclear epitopes but may disrupt membrane morphology.
AutofluorescenceIf background is high, reduce fixation time, improve washing, test a different blocking buffer, or switch fluorophores.

Tissue immunofluorescence protocol

Use this workflow for fluorescent antibody staining of frozen, fixed frozen, or FFPE tissue sections.

  • Use this format when: Detecting spatial protein expression in tissue sections with fluorescence rather than chromogenic IHC.
  • Optimize first: Section type, antigen retrieval, autofluorescence control, permeabilization, antibody dilution, and mounting medium.
1
Prepare sections
Use 4-8 um tissue sections mounted on charged slides. Air dry frozen sections for 10-20 minutes. For FFPE sections, deparaffinize through xylene and graded ethanol, then rehydrate to water.
2
Fix or retrieve antigen
For frozen sections, fix with cold acetone for 10 minutes or 4 percent paraformaldehyde for 10-15 minutes depending on the target. For FFPE sections, perform heat-induced antigen retrieval in citrate buffer pH 6.0 or Tris-EDTA pH 9.0 for 10-20 minutes, then cool 20 minutes.
3
Reduce autofluorescence if needed
For tissue with high autofluorescence, treat with an autofluorescence quenching reagent or freshly prepared 0.1 percent sodium borohydride for 5-10 minutes, followed by thorough PBS washes. Choose the method compatible with your fluorophores.
4
Permeabilize and block
Permeabilize with 0.1-0.3 percent Triton X-100 in PBS for 5-10 minutes if intracellular or nuclear access is needed. Block 30-60 minutes with 5 percent normal serum or 1-5 percent BSA in PBS.
5
Add primary antibody
Dilute ATOX1 Polyclonal Antibody in blocking buffer. Start with the datasheet-recommended tissue IF dilution or test 1:100, 1:250, and 1:500. Apply 100-200 uL per section, cover with parafilm or use a humidified chamber, and incubate 1-2 hours at room temperature or overnight at 4 degrees C.
6
Wash and add secondary antibody
Wash 3 times for 5 minutes each in PBS or PBST. Add fluorophore-conjugated secondary antibody diluted 1:500-1:1000. Incubate 45-60 minutes at room temperature protected from light.
7
Counterstain and mount
Wash 3 times. Add DAPI if desired. Mount with antifade medium and coverslip carefully to avoid bubbles.
8
Image sections
Use identical microscope settings for comparative samples. Include no-primary controls and known positive tissue when available.
Tissue handlingDo not let sections dry after rehydration or blocking begins.
RetrievalSome epitopes require retrieval for FFPE tissue but not frozen tissue. Test pH 6 and pH 9 retrieval when starting a new antibody.

Multiplex IF/ICC protocol

Use this workflow to detect two or more targets in the same cells or tissue section using spectrally distinct antibodies.

  • Use this format when: Comparing colocalization, marker identity, signaling state, or target expression in the same sample.
  • Optimize first: Host species compatibility, fluorophore separation, antibody order, blocking, cross-reactivity controls, and exposure settings.
1
Plan antibody panel
Choose primary antibodies from different host species when possible. Match secondary antibodies to non-overlapping fluorophores. Avoid fluorophores with heavy spectral overlap unless spectral unmixing is available.
2
Fix, permeabilize, and block
Prepare the sample using the IF/ICC or tissue IF fixation and blocking conditions appropriate for the most sensitive target. Use enough blocking buffer to reduce nonspecific secondary antibody binding.
3
Add primary antibody mix
Prepare a master mix containing each primary antibody at its optimized dilution. Include ATOX1 Polyclonal Antibody if it is one of the targets in the panel. Start with single-stain optimized dilutions before combining antibodies. Incubate 1 hour at room temperature or overnight at 4 degrees C.
4
Wash thoroughly
Wash 3 times for 5 minutes each with PBS or PBST. Extend washes to 5-10 minutes when background or cross-channel signal is high.
5
Add secondary antibody mix
Add cross-adsorbed secondary antibodies diluted in blocking buffer, commonly 1:500-1:1000. Incubate 45-60 minutes protected from light.
6
Counterstain and mount
Wash 3 times. Add nuclear counterstain if needed, then mount with antifade medium compatible with all fluorophores.
7
Acquire controls
Image single-color controls, no-primary controls, and full-stain samples using consistent settings. Use single-color controls to identify bleed-through or compensation needs.
Same-species primariesIf two primary antibodies share the same host species, use directly conjugated primaries, sequential staining with blocking, Fab fragments, or tyramide signal amplification when appropriate.
ColocalizationDo not call colocalization from overexposed images. Keep all channels below saturation.

Phospho/PTM IF protocol

Use this workflow for phosphorylation, acetylation, methylation, ubiquitination, or other modification-specific antibody staining.

  • Use this format when: Detecting transient signaling or post-translational modifications in cells or tissue.
  • Optimize first: Sample timing, inhibitor use, fixation speed, permeabilization, antibody dilution, and positive or negative treatment controls.
1
Prepare treated and control samples
Plan untreated, stimulated, inhibited, knockdown, or positive control samples before staining. For phospho targets, add phosphatase inhibitors during washes or lysis-related steps where applicable and fix quickly after treatment.
2
Fix rapidly
Remove medium and rinse briefly with warm PBS when needed. Fix with 4 percent paraformaldehyde for 10-15 minutes at room temperature, or use methanol fixation for targets known to work better with alcohol fixation.
3
Permeabilize for nuclear or intracellular access
Wash 3 times. Use 0.1-0.3 percent Triton X-100 for 5-10 minutes. For histone or chromatin PTMs, test 0.2-0.5 percent Triton X-100 and consider a short pre-extraction step only if compatible with morphology and the target.
4
Block
Block 45-60 minutes with 1-5 percent BSA in PBS or TBS. For phospho-specific antibodies, BSA is often preferred over milk because milk can contain phosphoproteins that increase background.
5
Add primary antibody
Dilute ATOX1 Polyclonal Antibody in blocking buffer. Start with the datasheet-recommended IF dilution or test 1:100, 1:250, and 1:500. Incubate overnight at 4 degrees C for low-abundance PTMs or weak signals.
6
Wash and detect
Wash 3 times for 5 minutes. Add fluorophore-conjugated secondary antibody at 1:500-1:1000 for 45-60 minutes protected from light. Wash 3 times.
7
Mount and image
Counterstain if needed, mount with antifade medium, and image positive and negative controls with the same settings. For signal quantification, avoid saturated pixels and analyze the same cellular compartment across samples.
PTM specificityUse inhibitor, stimulation, peptide competition, knockdown, or known positive/negative controls when available to support modification-specific interpretation.
Blocking bufferIf background is high, compare BSA, serum, fish gelatin, or commercial antibody diluent.
Open full IF/ICC protocol

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