I am now sharing an extremely detaild new method invented by the OpenAI GPT-5 Pro model for one of biology’s oldest and most foundational techniques: Western blotting. I had the honor of learning it in 1989 in Switzerland from two of its original inventors, Drs. Harry Towbin and Theo Staehelin.
This AI-generated scientific method aligns with what @kevinweil announced today about starting OpenAI for Science, an AI-powered platform aimed at accelerating scientific discovery. It also supports what I have long argued: models like GPT-5 Thinking and Pro can accelerate the scientific process by orders of magnitude; this is another example!
It is important for non-scientists to realize that technology drives science. Without novel scientific techniques and methods, we cannot sustain accelerated discovery. This is especially true in biology, which is extraordinarily complex, with trillions of parts and dynamically interacting processes. Without AI we would have little hope of understanding all this complexity and solving biology, curing all diseases, or reversing aging in the coming decades, or even centuries.
This is why I've always been so exicted about AI and went all in several years ago after ChatGPT release, when it became clear to me the path to AGI has been opened. It is now time to recognize that this is not a hypothetical future any more; it is already here and getting better by the month, week, and day!
I should also point out that further development and proof-of-concept experimentations could turn this particular method into a patentable invention. However, I strongly believe that in the age of AI, such inventions should not be patented and should belong to all of humanity. In fact, our patent and intellectual property laws need to be substantially modified, or even replaced, to align with the reality that soon most inventions may be generated by AI, with human contribution increasingly focused on application and value creation. I digress; that is a deeper topic for another time :)
Before I share the method developed by GPT-5 Pro in the following thread, I would like to provide some historical background and explain the importance of the Western blotting method, which every biologist knows or should know. Here I am sharing a summary of GPT-5’s explanation; of course, anyone can check with Grok or other models for more detail:
What is Western Blotting?
Think of a cell extract as a soup full of different proteins. Western blotting does four simple things:
Sorts by size. An electric field pulls proteins through a jelly-like slab. Small ones run farther than big ones.
Copies the pattern to paper. The separated proteins are moved onto a thin membrane that holds them in place.
Finds your protein with a matching “key.” An antibody that sticks only to your protein is added, then a second antibody with a built-in label makes the spot light up or change color. The position tells you the protein’s size, and the brightness tells you roughly how much is there.
Who invented it, and when?
In 1979, two groups independently created the core method of protein “blotting” to a membrane after gel electrophoresis: Harry Towbin, Theophil Staehelin, and Julian Gordon in Basel, and Jaime Renart, Jakob Reiser, and George Stark at Stanford. Two years later, in 1981, W. Neal Burnette popularized the name “western blot.”
Why is it a pillar of biological methods?
Historically and practically it is one of the core lab methods for protein analysis. The reason is its specificity with size information. It tells you not just that a protein is present, but also its approximate size, which helps confirm identity, isoforms, and post-translational modifications. Indeed, Western Blotting remains one of the most commonly used protein assays across research fields and has been central to countless studies and workflows. Labs use it to verify antibody targets, confirm expression or knockdowns, and check pathway activation and use it in diagnostics (for example early HIV testing was dependent on this).
I will share the detailed method in the next thread. It is important to point out that this is a very sophisticated method, that took several prompts to validate. However, it can continue to be improved and optimized further and of course significant lab effort will be needed to develop it and troubleshoot it. This is a very detailed blueprint, which in itself is extraordinarily remarkable! In the third thread below I provide some critiques (also from GPT-5 Pro) and how to further validate and further improve it.
Because this is very technical it may be quite difficult for those not familiar with these type of methods to follow, so here is a scientific and layperson’s summary of the gist:
DSI-Seq (novel western blotting replacement)
Scientific summary of DSI-Seq: Digital Size-Indexed ImmunoSequencing transforms the Western blot from an artisanal, low-plex picture into a standardized, high-plex, quantitative assay that preserves size-resolved proteoforms. By coupling rapid microchip SDS separation and inline fractionation to DNA-barcoded immunoassays with UMI-based counting, DSI-Seq delivers isoform-aware pathway readouts from tiny inputs with built-in controls and shareable digital outputs. This capability enables rigorous kinetic mapping of signaling, proteoform-level pharmacodynamic biomarkers, and reproducible QC for cell therapy and drug programs, filling a critical gap between legacy Westerns and high-bar mass spectrometry and advancing proteoform-centric biology across immunology and oncology.
Layperson summary: Think of DSI-Seq as a modern upgrade to the classic Western blot, the lab test that shows whether a protein is present and how big it is. Instead of a fuzzy picture of bands, it separates proteins by size on a tiny chip, adds simple barcodes, and then uses DNA reading tools to count each protein like a supermarket scanner. The result is a clear table of numbers that tells you not only how much of a protein is there, but also whether it is the full version, a cut piece, or a switched-on form. It can check many proteins at once from a very small sample and gives answers in hours instead of days. This helps scientists see how cells send signals, confirm how drugs and cell therapies work, and compare results across labs with much better consistency. In short, it turns a slow, manual art into a fast, reliable measurement that can speed up discoveries and improve testing in medicine.
This AI-generated scientific method aligns with what @kevinweil announced today about starting OpenAI for Science, an AI-powered platform aimed at accelerating scientific discovery. It also supports what I have long argued: models like GPT-5 Thinking and Pro can accelerate the scientific process by orders of magnitude; this is another example!
It is important for non-scientists to realize that technology drives science. Without novel scientific techniques and methods, we cannot sustain accelerated discovery. This is especially true in biology, which is extraordinarily complex, with trillions of parts and dynamically interacting processes. Without AI we would have little hope of understanding all this complexity and solving biology, curing all diseases, or reversing aging in the coming decades, or even centuries.
This is why I've always been so exicted about AI and went all in several years ago after ChatGPT release, when it became clear to me the path to AGI has been opened. It is now time to recognize that this is not a hypothetical future any more; it is already here and getting better by the month, week, and day!
I should also point out that further development and proof-of-concept experimentations could turn this particular method into a patentable invention. However, I strongly believe that in the age of AI, such inventions should not be patented and should belong to all of humanity. In fact, our patent and intellectual property laws need to be substantially modified, or even replaced, to align with the reality that soon most inventions may be generated by AI, with human contribution increasingly focused on application and value creation. I digress; that is a deeper topic for another time :)
Before I share the method developed by GPT-5 Pro in the following thread, I would like to provide some historical background and explain the importance of the Western blotting method, which every biologist knows or should know. Here I am sharing a summary of GPT-5’s explanation; of course, anyone can check with Grok or other models for more detail:
What is Western Blotting?
Think of a cell extract as a soup full of different proteins. Western blotting does four simple things:
Sorts by size. An electric field pulls proteins through a jelly-like slab. Small ones run farther than big ones.
Copies the pattern to paper. The separated proteins are moved onto a thin membrane that holds them in place.
Finds your protein with a matching “key.” An antibody that sticks only to your protein is added, then a second antibody with a built-in label makes the spot light up or change color. The position tells you the protein’s size, and the brightness tells you roughly how much is there.
Who invented it, and when?
In 1979, two groups independently created the core method of protein “blotting” to a membrane after gel electrophoresis: Harry Towbin, Theophil Staehelin, and Julian Gordon in Basel, and Jaime Renart, Jakob Reiser, and George Stark at Stanford. Two years later, in 1981, W. Neal Burnette popularized the name “western blot.”
Why is it a pillar of biological methods?
Historically and practically it is one of the core lab methods for protein analysis. The reason is its specificity with size information. It tells you not just that a protein is present, but also its approximate size, which helps confirm identity, isoforms, and post-translational modifications. Indeed, Western Blotting remains one of the most commonly used protein assays across research fields and has been central to countless studies and workflows. Labs use it to verify antibody targets, confirm expression or knockdowns, and check pathway activation and use it in diagnostics (for example early HIV testing was dependent on this).
I will share the detailed method in the next thread. It is important to point out that this is a very sophisticated method, that took several prompts to validate. However, it can continue to be improved and optimized further and of course significant lab effort will be needed to develop it and troubleshoot it. This is a very detailed blueprint, which in itself is extraordinarily remarkable! In the third thread below I provide some critiques (also from GPT-5 Pro) and how to further validate and further improve it.
Because this is very technical it may be quite difficult for those not familiar with these type of methods to follow, so here is a scientific and layperson’s summary of the gist:
DSI-Seq (novel western blotting replacement)
Scientific summary of DSI-Seq: Digital Size-Indexed ImmunoSequencing transforms the Western blot from an artisanal, low-plex picture into a standardized, high-plex, quantitative assay that preserves size-resolved proteoforms. By coupling rapid microchip SDS separation and inline fractionation to DNA-barcoded immunoassays with UMI-based counting, DSI-Seq delivers isoform-aware pathway readouts from tiny inputs with built-in controls and shareable digital outputs. This capability enables rigorous kinetic mapping of signaling, proteoform-level pharmacodynamic biomarkers, and reproducible QC for cell therapy and drug programs, filling a critical gap between legacy Westerns and high-bar mass spectrometry and advancing proteoform-centric biology across immunology and oncology.
Layperson summary: Think of DSI-Seq as a modern upgrade to the classic Western blot, the lab test that shows whether a protein is present and how big it is. Instead of a fuzzy picture of bands, it separates proteins by size on a tiny chip, adds simple barcodes, and then uses DNA reading tools to count each protein like a supermarket scanner. The result is a clear table of numbers that tells you not only how much of a protein is there, but also whether it is the full version, a cut piece, or a switched-on form. It can check many proteins at once from a very small sample and gives answers in hours instead of days. This helps scientists see how cells send signals, confirm how drugs and cell therapies work, and compare results across labs with much better consistency. In short, it turns a slow, manual art into a fast, reliable measurement that can speed up discoveries and improve testing in medicine.
Method: Digital Size‑Indexed ImmunoSequencing (DSI‑Seq)
Goal: preserve Western’s size information and antibody specificity, but make it quantitative, multiplexed, fast, and low input.
Core idea
Rapid microchip SDS separation of denatured proteins.
Inline fractionation of the eluting stream into many narrow size bins.
Multiplex DNA‑barcoded immunoassay in each bin.
Sequencing or digital PCR readout to count molecules per target per size bin.
Computational mapping from bin index to molecular weight using co‑run standards.
This yields a 2D matrix: targets on one axis, size bins on the other. You see isoforms and shifts exactly like a blot, but with digital counts instead of gray bands.
Why it beats Westerns
Isoform-resolved, like a blot: size bins let you separate full-length vs cleaved or shifted proteoforms.
High multiplex: 50 to 200 targets per run is realistic with barcoded antibodies.
Quantitative: counts are digital, with internal spike‑ins and UMIs for normalization and linearity.
Low sample: nanoliter fractions and single‑tube chemistry cut input requirements.
Throughput: dozens of samples per day on a benchtop box, no membranes, no overnight incubations.
Reproducibility: fixed microfluidic geometry, DNA barcodes with error correction, built‑in standards.
Workflow overview
Sample prep
Lyse in SDS buffer with protease and phosphatase inhibitors as needed.
Add a small panel of recombinant protein ladder standards spanning 10 to 250 kDa.
Microchip SDS separation
Use a disposable polymer‑sieving microfluidic chip.
Inject sample plus ladder. Separate over a short channel. Total runtime on the order of minutes.
Inline fractionation
At the channel outlet, segment the eluent into nanoliter droplets at a fixed frequency.
Each droplet receives a time‑stamp DNA barcode from a clocked side stream. The time stamp uniquely encodes the size bin because migration time maps to molecular weight. In plate‑based builds, collect sequential fractions across 64 to 128 wells instead of droplets and dispense a unique DNA bin barcode into each well.
Multiplex immunoassay with DNA barcodes
Add a cocktail of antibodies, each conjugated to a unique DNA tag that carries an antibody ID barcode, a unique molecular identifier (UMI), universal priming sites.
Two options for specificity:
Single‑epitope capture: antibodies on magnetic beads capture target in each bin. A reporter oligo is released only upon capture using a proximity‑restricted cleavage or displacement.
Proximity extension assay (PEA): two oligo‑tagged antibodies per target give a ligation‑competent DNA only when both bind the same protein. This strongly reduces off‑target.
The bin time‑stamp barcode is ligated to each target amplicon so every read is labeled with both protein identity and size bin.
Readout
PCR amplify and pool. Quantify by NGS for high plex or by multiplex digital PCR for 5 to 30 targets.
Demultiplex reads to counts per target per bin. Collapse UMIs to remove amplification bias.
Analysis
Fit ladder‑derived calibration to convert bin index to apparent kDa.
Call peaks per target to quantify proteoforms or shifts.
Normalize using: external spike‑in protein standards, per‑bin total protein dye signal, or stable reference proteins.
You can plot fraction profiles the way you read bands now, but with real numbers and confidence intervals.
Key design details
Microfluidics
Short, linear polyacrylamide sieving matrix in a molded chip.
64 to 128 fraction bins across the separation window. Effective resolution comparable to 6 to 12 percent gels.
Droplet mode or 96‑well fractionation module, depending on comfort level.
Barcodes and error control
12 to 16 nt antibody ID barcodes with Hamming distance of 3 or more.
8 to 12 nt time‑stamp barcodes for bins.
8 to 12 nt UMIs to deduplicate PCR bias.
Use a universal primer pair for all targets to simplify amplification.
Antibody conjugation
Protein A/G mediated orientation, then NHS‑PEG‑azide to click a maleimide‑bearing oligo, or use site‑specific conjugation on engineered Fc.
Validate each antibody’s linear epitope binding under 0.05 to 0.1 percent SDS or after SDS quench with cyclodextrin.
Controls
No‑antibody and isotype controls in a few bins to estimate background.
Known‑ratio mixtures of recombinant targets to test linearity.
Phosphatase‑treated lysate as negative control for phospho‑specific panels.
Performance targets to aim for
Multiplex: 100 proteins per reaction with NGS; 10 to 30 with dPCR.
Dynamic range: 4 to 5 orders of magnitude with UMIs and spike‑ins.
Sensitivity: low femtomoles per target per lane is realistic; sub‑microgram total protein per sample.
Size resolution: 5 to 10 percent across 15 to 250 kDa with 64 to 128 bins.
Hands‑on time: minimal, no membranes or overnight incubations.
These are engineering targets, not guarantees. They are chosen based on what each subsystem can credibly deliver.
Validation plan that will convince a hard skeptic
Build the separation plus fractionation stub
Run a prestained ladder through the chip.
Fractionate into 64 bins.
Confirm bin‑to‑kDa mapping with a logistic fit.
Single‑target end‑to‑end test
One antibody with an oligo tag.
Spike recombinant protein at a dilution series into a HeLa lysate.
Show linear counts versus input and a single bin peak at the expected kDa.
Isoform resolution
PARP1 cleavage in apoptosis: treat cells with staurosporine.
Detect 116 kDa full length and 89 kDa cleaved in separate bins.
PTM specificity
Phospho‑ERK1/2: stimulate and phosphatase‑treat controls.
Phospho‑specific antibody should yield bin‑aligned signal that collapses with phosphatase.
Test Multiplex panel
24‑plex T cell signaling panel: CD3ζ pY, ZAP70 pY, LAT pY, SLP76, PLCG1 pY, ERK1/2 pT/pY, AKT pS473, mTOR pS2448, 4EBP1 pT37/46, NF‑κB p65 pS536, beta‑actin as reference.
Compare fold changes to traditional Westerns and phospho‑flow as orthogonal methods.
Reproducibility
10 technical replicates. Compute CV per target per bin.
Batch‑to‑batch chip variation and antibody lot testing.
Practical build choices
Version A: plate‑basedUse capillary electrophoresis into a 96‑well plate, dispense pre‑aliquoted bin barcodes by row, perform the immuno‑DNA chemistry in wells, and read by amplicon sequencing. This is the fastest route with standard gear.
Version B: droplet‑basedMicrofluidic droplet generator assigns the bin time‑stamp barcode in flow, merges with antibody beads, then breaks emulsion for pooled amplification. Higher automation and less loss, but more microfluidic engineering.
Version C: no‑sequencer optionReplace NGS with a panel of TaqMan probes on a digital PCR platform. Lower plex but no dependence on sequencing.
Risks and mitigations
Antibody performance in residual SDSMitigation: dilute SDS below 0.05 percent, use SDS scavengers, favor linear epitope antibodies, or renature briefly before binding.
Carryover between binsMitigation: increase droplet segmentation rate or fraction dead volumes, add a short waste gap between bins, validate cross‑talk with ladder only.
Barcode cross‑talk and index hoppingMitigation: error‑correcting barcodes, unique dual‑indexing, UMI deduplication.
Off‑target binding in complex lysatesMitigation: use PEA two‑antibody logic for problematic targets, include matched isotype and competition controls.
Mapping accuracy from time to kDaMitigation: co‑run ladder and fit per run. Report apparent kDa with confidence intervals.
How it compares to current alternatives
Traditional Western: size info yes, multiplex low, quantitation poor, time long. DSI‑Seq: size info yes, multiplex high, digital counts, faster.
Capillary Western systems: automated and quantitative but limited multiplex. DSI‑Seq: retains automation but scales multiplex via barcodes.
DigiWest‑like bead fractionation: conceptually close but uses protein‑on‑bead plus fluorescent detection. DSI‑Seq: swaps fluorescence for DNA counting with UMIs and a simpler microchip separation.
Targeted mass spectrometry: high specificity and absolute quantitation, but complex setup and limited size visualization. DSI‑Seq: more accessible and preserves a blot‑like picture of proteoforms.
Kit and instrument concept
Disposable chip with sieving matrix and outlet fractionator.
Antibody panel with validated DNA barcodes and UMIs.
Bin barcode plate or droplet side‑stream mix pre‑made.
Calibration ladder proteins plus spike‑in counting controls.
Benchtop controller for voltage, flows, and droplet timing.
Software that auto‑maps bins to kDa, calls peaks, and outputs publication‑ready plots and CSVs.
When denaturation or SDS is unavoidable
If you must process denatured lysate, dilute or scavenge SDS before proximity chemistry. Many Western‑validated antibodies bind linear epitopes after SDS quench.
If two‑epitope binding fails under those conditions, switch the target to Option B capture‑and‑release for that protein while keeping proximity extension for the rest.
Failure modes to plan for, with fixes
High background without protein: incomplete nuclease inactivation or oligo self‑ligation. Fix by tightening the nuclease step and redesigning oligos with higher Hamming distance and blocked ends.
Hook effect at very high abundance: split samples or dilute to keep partition occupancy between 0.1 and 0.7.
Poor agreement between targets: recalibrate conversion efficiencies with purified standards, and check antibody pair compatibility.
Cross‑reactivity: migrate problematic targets to the capture‑and‑release format or require a competition control for acceptance.
Bottom line and cross-check with the landscape:
DSI‑Seq is a novel and feasible direction for a true “Western‑replacement” that preserves size information while giving digital, multiplexed readouts. The closest prior arts solve only parts of this: DigiWest multiplexes after SDS‑PAGE but uses bead fluorescence, not sequencing; Simple Western automates size‑resolved immunoassays but does not scale to high plex; Olink PEA and other DNA‑barcoded immunoassays are highly multiplexed but lack size resolution. Combination of microchip SDS separation + inline fractionation + bin barcoding + DNA‑counting does not appear in the literature as a unified method.
How it compares to what exists
DigiWest: SDS‑PAGE lane is sliced and eluted to barcoded beads, then probed and read by Luminex fluorescence. It retains size info and multiplexes, but detection is not digital sequencing. Your approach swaps in sequencing with UMIs and a microchip separation front‑end.
Simple Western (capillary Western): automated, quantitative, but limited multiplex and no sequencing readout.
DNA‑barcoded immunoassays: Olink PEA and ID‑seq give high plex digital counts by NGS, but they operate in solution without size resolution. You add back the Western’s unique size axis.
Microchip SDS protein sizing: commercially standard with typical ∼10 percent sizing resolution in minutes, which matches your binning targets.
Conclusion on novelty: the integration and the bin time‑stamping idea look patentable and practically differentiating.
Goal: preserve Western’s size information and antibody specificity, but make it quantitative, multiplexed, fast, and low input.
Core idea
Rapid microchip SDS separation of denatured proteins.
Inline fractionation of the eluting stream into many narrow size bins.
Multiplex DNA‑barcoded immunoassay in each bin.
Sequencing or digital PCR readout to count molecules per target per size bin.
Computational mapping from bin index to molecular weight using co‑run standards.
This yields a 2D matrix: targets on one axis, size bins on the other. You see isoforms and shifts exactly like a blot, but with digital counts instead of gray bands.
Why it beats Westerns
Isoform-resolved, like a blot: size bins let you separate full-length vs cleaved or shifted proteoforms.
High multiplex: 50 to 200 targets per run is realistic with barcoded antibodies.
Quantitative: counts are digital, with internal spike‑ins and UMIs for normalization and linearity.
Low sample: nanoliter fractions and single‑tube chemistry cut input requirements.
Throughput: dozens of samples per day on a benchtop box, no membranes, no overnight incubations.
Reproducibility: fixed microfluidic geometry, DNA barcodes with error correction, built‑in standards.
Workflow overview
Sample prep
Lyse in SDS buffer with protease and phosphatase inhibitors as needed.
Add a small panel of recombinant protein ladder standards spanning 10 to 250 kDa.
Microchip SDS separation
Use a disposable polymer‑sieving microfluidic chip.
Inject sample plus ladder. Separate over a short channel. Total runtime on the order of minutes.
Inline fractionation
At the channel outlet, segment the eluent into nanoliter droplets at a fixed frequency.
Each droplet receives a time‑stamp DNA barcode from a clocked side stream. The time stamp uniquely encodes the size bin because migration time maps to molecular weight. In plate‑based builds, collect sequential fractions across 64 to 128 wells instead of droplets and dispense a unique DNA bin barcode into each well.
Multiplex immunoassay with DNA barcodes
Add a cocktail of antibodies, each conjugated to a unique DNA tag that carries an antibody ID barcode, a unique molecular identifier (UMI), universal priming sites.
Two options for specificity:
Single‑epitope capture: antibodies on magnetic beads capture target in each bin. A reporter oligo is released only upon capture using a proximity‑restricted cleavage or displacement.
Proximity extension assay (PEA): two oligo‑tagged antibodies per target give a ligation‑competent DNA only when both bind the same protein. This strongly reduces off‑target.
The bin time‑stamp barcode is ligated to each target amplicon so every read is labeled with both protein identity and size bin.
Readout
PCR amplify and pool. Quantify by NGS for high plex or by multiplex digital PCR for 5 to 30 targets.
Demultiplex reads to counts per target per bin. Collapse UMIs to remove amplification bias.
Analysis
Fit ladder‑derived calibration to convert bin index to apparent kDa.
Call peaks per target to quantify proteoforms or shifts.
Normalize using: external spike‑in protein standards, per‑bin total protein dye signal, or stable reference proteins.
You can plot fraction profiles the way you read bands now, but with real numbers and confidence intervals.
Key design details
Microfluidics
Short, linear polyacrylamide sieving matrix in a molded chip.
64 to 128 fraction bins across the separation window. Effective resolution comparable to 6 to 12 percent gels.
Droplet mode or 96‑well fractionation module, depending on comfort level.
Barcodes and error control
12 to 16 nt antibody ID barcodes with Hamming distance of 3 or more.
8 to 12 nt time‑stamp barcodes for bins.
8 to 12 nt UMIs to deduplicate PCR bias.
Use a universal primer pair for all targets to simplify amplification.
Antibody conjugation
Protein A/G mediated orientation, then NHS‑PEG‑azide to click a maleimide‑bearing oligo, or use site‑specific conjugation on engineered Fc.
Validate each antibody’s linear epitope binding under 0.05 to 0.1 percent SDS or after SDS quench with cyclodextrin.
Controls
No‑antibody and isotype controls in a few bins to estimate background.
Known‑ratio mixtures of recombinant targets to test linearity.
Phosphatase‑treated lysate as negative control for phospho‑specific panels.
Performance targets to aim for
Multiplex: 100 proteins per reaction with NGS; 10 to 30 with dPCR.
Dynamic range: 4 to 5 orders of magnitude with UMIs and spike‑ins.
Sensitivity: low femtomoles per target per lane is realistic; sub‑microgram total protein per sample.
Size resolution: 5 to 10 percent across 15 to 250 kDa with 64 to 128 bins.
Hands‑on time: minimal, no membranes or overnight incubations.
These are engineering targets, not guarantees. They are chosen based on what each subsystem can credibly deliver.
Validation plan that will convince a hard skeptic
Build the separation plus fractionation stub
Run a prestained ladder through the chip.
Fractionate into 64 bins.
Confirm bin‑to‑kDa mapping with a logistic fit.
Single‑target end‑to‑end test
One antibody with an oligo tag.
Spike recombinant protein at a dilution series into a HeLa lysate.
Show linear counts versus input and a single bin peak at the expected kDa.
Isoform resolution
PARP1 cleavage in apoptosis: treat cells with staurosporine.
Detect 116 kDa full length and 89 kDa cleaved in separate bins.
PTM specificity
Phospho‑ERK1/2: stimulate and phosphatase‑treat controls.
Phospho‑specific antibody should yield bin‑aligned signal that collapses with phosphatase.
Test Multiplex panel
24‑plex T cell signaling panel: CD3ζ pY, ZAP70 pY, LAT pY, SLP76, PLCG1 pY, ERK1/2 pT/pY, AKT pS473, mTOR pS2448, 4EBP1 pT37/46, NF‑κB p65 pS536, beta‑actin as reference.
Compare fold changes to traditional Westerns and phospho‑flow as orthogonal methods.
Reproducibility
10 technical replicates. Compute CV per target per bin.
Batch‑to‑batch chip variation and antibody lot testing.
Practical build choices
Version A: plate‑basedUse capillary electrophoresis into a 96‑well plate, dispense pre‑aliquoted bin barcodes by row, perform the immuno‑DNA chemistry in wells, and read by amplicon sequencing. This is the fastest route with standard gear.
Version B: droplet‑basedMicrofluidic droplet generator assigns the bin time‑stamp barcode in flow, merges with antibody beads, then breaks emulsion for pooled amplification. Higher automation and less loss, but more microfluidic engineering.
Version C: no‑sequencer optionReplace NGS with a panel of TaqMan probes on a digital PCR platform. Lower plex but no dependence on sequencing.
Risks and mitigations
Antibody performance in residual SDSMitigation: dilute SDS below 0.05 percent, use SDS scavengers, favor linear epitope antibodies, or renature briefly before binding.
Carryover between binsMitigation: increase droplet segmentation rate or fraction dead volumes, add a short waste gap between bins, validate cross‑talk with ladder only.
Barcode cross‑talk and index hoppingMitigation: error‑correcting barcodes, unique dual‑indexing, UMI deduplication.
Off‑target binding in complex lysatesMitigation: use PEA two‑antibody logic for problematic targets, include matched isotype and competition controls.
Mapping accuracy from time to kDaMitigation: co‑run ladder and fit per run. Report apparent kDa with confidence intervals.
How it compares to current alternatives
Traditional Western: size info yes, multiplex low, quantitation poor, time long. DSI‑Seq: size info yes, multiplex high, digital counts, faster.
Capillary Western systems: automated and quantitative but limited multiplex. DSI‑Seq: retains automation but scales multiplex via barcodes.
DigiWest‑like bead fractionation: conceptually close but uses protein‑on‑bead plus fluorescent detection. DSI‑Seq: swaps fluorescence for DNA counting with UMIs and a simpler microchip separation.
Targeted mass spectrometry: high specificity and absolute quantitation, but complex setup and limited size visualization. DSI‑Seq: more accessible and preserves a blot‑like picture of proteoforms.
Kit and instrument concept
Disposable chip with sieving matrix and outlet fractionator.
Antibody panel with validated DNA barcodes and UMIs.
Bin barcode plate or droplet side‑stream mix pre‑made.
Calibration ladder proteins plus spike‑in counting controls.
Benchtop controller for voltage, flows, and droplet timing.
Software that auto‑maps bins to kDa, calls peaks, and outputs publication‑ready plots and CSVs.
When denaturation or SDS is unavoidable
If you must process denatured lysate, dilute or scavenge SDS before proximity chemistry. Many Western‑validated antibodies bind linear epitopes after SDS quench.
If two‑epitope binding fails under those conditions, switch the target to Option B capture‑and‑release for that protein while keeping proximity extension for the rest.
Failure modes to plan for, with fixes
High background without protein: incomplete nuclease inactivation or oligo self‑ligation. Fix by tightening the nuclease step and redesigning oligos with higher Hamming distance and blocked ends.
Hook effect at very high abundance: split samples or dilute to keep partition occupancy between 0.1 and 0.7.
Poor agreement between targets: recalibrate conversion efficiencies with purified standards, and check antibody pair compatibility.
Cross‑reactivity: migrate problematic targets to the capture‑and‑release format or require a competition control for acceptance.
Bottom line and cross-check with the landscape:
DSI‑Seq is a novel and feasible direction for a true “Western‑replacement” that preserves size information while giving digital, multiplexed readouts. The closest prior arts solve only parts of this: DigiWest multiplexes after SDS‑PAGE but uses bead fluorescence, not sequencing; Simple Western automates size‑resolved immunoassays but does not scale to high plex; Olink PEA and other DNA‑barcoded immunoassays are highly multiplexed but lack size resolution. Combination of microchip SDS separation + inline fractionation + bin barcoding + DNA‑counting does not appear in the literature as a unified method.
How it compares to what exists
DigiWest: SDS‑PAGE lane is sliced and eluted to barcoded beads, then probed and read by Luminex fluorescence. It retains size info and multiplexes, but detection is not digital sequencing. Your approach swaps in sequencing with UMIs and a microchip separation front‑end.
Simple Western (capillary Western): automated, quantitative, but limited multiplex and no sequencing readout.
DNA‑barcoded immunoassays: Olink PEA and ID‑seq give high plex digital counts by NGS, but they operate in solution without size resolution. You add back the Western’s unique size axis.
Microchip SDS protein sizing: commercially standard with typical ∼10 percent sizing resolution in minutes, which matches your binning targets.
Conclusion on novelty: the integration and the bin time‑stamping idea look patentable and practically differentiating.
Here is the interrogation of feasibility, potential issues problems, how to solve them, improvements to the assay, quality checks, go no go acceptance criteria:
Feasibility verdict
Technically feasible with careful chemistry and controls. The gating risk is background signal from oligo interactions and matrix inhibition of the polymerase. The second risk is biology specific, namely finding antibody pairs that both bind the same protein in lysate while tolerating the buffer you need. Both can be engineered down to acceptable levels with the mitigation stack below.
If your goal is accurate copy numbers for 10 to 20 proteins per sample in standard lysates, this can be achieved. For membrane proteins or harsh buffers, plan to use a bead capture step to remove inhibitors before PCR.
Key risks and how to retire them fast
Antibody binding after SDS Go/no‑go: binding at ≤0.01 percent SDS after cyclodextrin treatment must recover to at least 60–80 percent of native control for a test panel. If not, switch to a rapid buffer‑exchange or on‑bead SDS removal before binding.
Bin cross‑talk Quantify with a fluorescent ladder only. Require <3 percent signal in adjacent bins for the dominant band before moving on.
Barcode cross‑talk and index hopping Use unique dual indexes with error‑correcting antibody ID barcodes. Enforce strict adapter cleanup.
Insufficient effective resolution If dispersion broadens peaks across >3 bins at mid‑range, increase number of bins and shorten the separation window. Note that commercial chips deliver ∼10 percent resolution, which maps well to 64–128 bins over 15–250 kDa.
UMI collisions at high copy bins Use 14–16 nt UMIs and UMI‑aware deduplication tools.
Recommended MVP path
Prioritize Version A, plate‑based. It is the fastest credible build with standard gear.
Step 1. Build separation + fractionation stub
Microchip SDS run into a 96‑well plate. Pre‑aliquot bin barcodes across wells. Confirm bin‑to‑kDa calibration with a prestained ladder.
Acceptance: logistic fit r² ≥ 0.98, sizing error ≤ 10 percent for ladder bands.
Step 2. SDS neutralization check
Titrate β‑cyclodextrin in collected fractions and test one antibody–antigen pair known to be sensitive to SDS. Require ≥ 60 percent of native binding at ≤ 0.01 percent residual SDS.
Step 3. Single‑target end‑to‑end
One DNA‑tagged antibody, HeLa lysate spiked with recombinant target, 8‑point dilution, NGS readout with 16 nt UMIs. Expect linear counts over at least 3 logs with UMI collapse.
Step 4. Isoform resolution
PARP1 cleavage model. Show two non‑overlapping peaks at ∼116 and ∼89 kDa with <10 percent cross‑bin bleed.
Step 5. PTM specificity
ERK1/2 pT/pY with phosphatase control. Phospho signal should vanish in treated samples while total ERK remains. Use PEA for phospho to suppress off‑targets.
Step 6. 24‑plex T‑cell panel
Run your listed phospho panel. Benchmark fold‑changes against Simple Western and phospho‑flow. Expect rank‑order agreement and tighter CVs on DSI‑Seq if normalization is done with spike‑ins and per‑bin protein signal.
Quantitative design notes
Bins and segmentation: with a 2–3 minute separation window and 96–128 bins, you are only asking for ∼0.5–1.0 Hz effective binning. That is easy for a valve or droplet generator, and nanoliter per bin volumes are compatible with standard PCR chemistry. Reviews document CE‑to‑droplet interfaces.
Normalization hierarchy: 1) external spike‑in protein standards, 2) per‑bin total protein measurement via a dye read on the plate before PCR, 3) stable reference proteins.
Library design: error‑correcting 12–16 nt antibody IDs with Hamming distance ≥ 3, 14–16 nt UMIs, and unique dual indexes for samples.
Data analysis: UMI collapse, per‑bin background modeling, then peak calling on smoothed target‑wise bin profiles with FDR control. UMI‑aware tools will handle sequencing errors in UMIs.
Where to be extra hard on yourselves
SDS compatibility is the make‑or‑break. Literature is clear that SDS suppresses immunorecognition in solution. Beta‑cyclodextrin removal is credible, but you must prove quantitative recovery for several antibody classes and for phospho‑epitopes.
True size fidelity. Demand bin‑to‑kDa errors ≤ 10 percent across the ladder. Commercial microchip SDS assays hit this spec, so you should too.
Cross‑method agreement. In the 24‑plex test, require high concordance with Simple Western and targeted MS for several anchors. Targeted MS is the recognized Western alternative for quantitative work. Use it to certify your dynamic range and linearity.
Hard problems you must solve, with fixes
One protein should create one amplicon
Risk: proximity oligos extend or ligate without protein, or multivalent proteins give more than one product.
Fixes:
Use two oligos that are each locked in a short hairpin. A short connector strand opens both hairpins only when co‑localized on the same protein, then a high‑fidelity polymerase fills in to create the amplicon. Include 3′‑phosphate blocking until the connector is present.
For oligomers, design epitope pairs on a single monomer when possible. If not possible, report results per complex rather than per monomer and state that explicitly.
Antibody pair availability and behavior in lysate
Risk: lack of two non‑overlapping linear epitopes, steric clash, or loss of binding in detergent.
Fixes:
Start with clones validated for Western or IP, not only IF. Western and IP clones tolerate linear epitopes and exposure to detergents.
Perform epitope binning by BLI or SPR to choose non‑competing pairs separated by at least 5 to 10 nm on the unfolded or partially refolded protein model.
Where pairs are weak, replace one antibody with a nanobody or aptamer against a distinct linear epitope.
For phospho targets, prefer two antibodies that bind outside the modified residue, with one being modification specific. If this is impossible, move that target to capture‑and‑release.
Matrix inhibition of the proximity and PCR steps
Risk: detergents, salts, or protease inhibitors compromise extension and PCR.
Fixes:
Keep lysis buffer PCR compatible when possible. If you must use strong detergents, run capture‑and‑release: bead capture, wash to remove inhibitors, then release DNA reporter into clean PCR buffer.
Add a short post‑binding buffer exchange on magnetic beads for problematic targets.
Validate a polymerase mix that tolerates residual lysate at your intended dilution.
Background from carryover DNA or oligo self‑assembly
Risk: false positives dominate the low end and inflate the apparent copy number.
Fixes:
Pre‑clear the lysate with Benzonase, then inactivate it completely before proximity chemistry.
Use dUTP in all amplicons and add UDG before PCR to destroy any carryover products from previous runs.
Split pre‑PCR and post‑PCR spaces physically, with aerosol‑barrier tips and separate lab coats.
Sequence design: hairpin locks, high Hamming distance between barcodes, blocked ends, and no complementarity between any non‑partner oligos. Reject any panel where the reagent‑only control exceeds 0.2 positive partitions per 20,000 partitions.
Hook effect and nonlinearity at high abundance
Risk: signal per protein falls at high concentration because the two binders distribute on different molecules.
Fixes:
Titrate antibody concentrations into the zone where both epitopes are saturated for your expected protein range.
Always include a two‑point dilution series per sample for the abundant proteins. Accept a target only if the two dilutions agree within 10 percent after Poisson correction.
Multiplex cross‑talk
Risk: oligos from different pairs hybridize or amplify off target.
Fixes:
Build the panel by subpanels. Screen each new pair against the existing set for off‑target positives in a protein‑free background.
If a pair misbehaves, move it to the capture‑and‑release lane to isolate its chemistry.
Go or no‑go acceptance criteria
Adopt these hard thresholds before you believe a number.
Reagent‑only background
Less than 0.2 positives per 20,000 partitions per target channel.
Blank lysate after nuclease
Less than 0.5 positives per 20,000 partitions with universal primers but without proximity chemistry. If not, your nuclease or physical separation is insufficient.
Spike‑in recovery
Recovery 0.8 to 1.2 across a 100‑fold range for purified protein spiked into matrix. Failures trigger re‑titration of antibody concentrations and buffer cleanup.
Dilution linearity in real lysate
Two dilutions that differ by 2× must report a 1.8 to 2.2 ratio after Poisson correction.
Repeatability
Technical replicate CV less than 10 percent at 400 to 10,000 molecules per reaction.
Cross‑panel interference
Adding or removing other antibody pairs changes the measured copy number by less than 10 percent.
Bead‑gated proximityBind the capture antibody on beads, add the reporter antibody in solution, perform proximity extension on‑bead, then wash and elute the newly formed DNA into PCR buffer. This removes inhibitors and unreacted oligos and cuts background sharply. It reduces hands‑off simplicity but pays off for tough targets.
Connector‑mediated extension with dual locksEncode two short toeholds on the oligo arms and design a connector that bridges both with perfect Tm only when both arms are present. Include 3′ phosphates on arms and a 5′ phosphate on the connector. Use a strand‑displacing polymerase that starts only after ligation or nick‑translation. This reduces unspecific arm‑to‑arm pairing.
UDG carryover suppression and separate primer poolsUse dUTP in all products plus UDG in the PCR master mix. Maintain separate primer mixes for each subpanel to avoid building a universal contaminant that can amplify anywhere.
External protein counting standardInclude a recombinant protein with an artificial two‑epitope tag and its dedicated antibody pair in every reaction at a fixed copy number. This provides an internal process control for conversion efficiency and partition calling. Report all targets relative to this control and then convert back to absolute units with its known copy number.
Per‑target conversion efficiency calibrationMeasure epsilon for each target with a dilution series of purified protein in matrix. Store epsilon in software and apply a simple correction: estimated molecules equal measured molecules divided by epsilon. Update epsilon with each new antibody lot.
Panel design discipline
Keep amplicon length between 60 and 100 nt with matched Tm, design all probes against non‑overlapping internal sequences, and require Hamming distance of at least 3 for barcodes. Reject any oligo that creates primer dimers in silico against the entire panel.
PTM‑aware strategiesFor phosphorylation, when two antibodies cannot both bind, pair a pan antibody with a phospho‑specific binder and place the oligo on the pan arm only. Then use capture‑and‑release where phospho recognition triggers DNA release. This keeps the readout phospho dependent without needing both arms to bind.
Membrane protein handlingFor multi‑pass proteins, use mild detergents such as digitonin or DDM for extraction, then switch to bead capture with washes to remove detergents before the proximity step.
Failure modes you will see first, and how to debug in one day
Everything is positive
Likely contamination with prior amplicon. Replace all solutions, add UDG, and move pre‑ and post‑PCR physically apart. Confirm by running reagent‑only controls.
Nothing is positive, even with recombinant protein
Connector or polymerase step is not working, or one epitope is masked. Verify each arm can be extended alone with a synthetic splint, check antibody binding by classic sandwich ELISA, then return to proximity chemistry.
Strong signal in no‑protein control
Your arms are self‑complementary or the connector bridges unrelated arms. Redesign oligos with hairpins and toeholds that require co‑localization.
Nonlinear response across dilutions
You are in hook territory or conversion efficiency varies with protein concentration. Lower antibody concentration or split the sample. Re‑titrate to achieve flat efficiency across the intended range.
Statistics you should report with every result
Molecules per reaction with 95 percent confidence intervals based on binomial uncertainty of positives, converted via the standard Poisson occupancy model.
Dilution agreement score for each target.
Conversion efficiency epsilon and its uncertainty if you apply the correction.
Background positives per 20,000 partitions for the run.
Practical build choices I recommend
Start plate‑based. Use a microchip SDS instrument or capillary interface to a 96‑well plate, pre‑aliquot bin barcodes, do all chemistry in wells. This gets you data with minimal custom microfluidics. Published interfaces from electrophoresis to droplets or plates give you patterns to copy.
PEA for phospho targets, single‑epitope capture for abundant structural targets. PEA is proven to reduce off‑target noise in multiplex settings.
SDS removal: add β‑cyclodextrin and a brief dilution step upon fraction entry, then bind. Validate residual SDS by a colorimetric assay or mass‑balance against standards.
Controls: isotypes, no‑antibody bins, phosphatase controls for phospho, and known‑ratio recombinant mixes to certify linearity. This mirrors best practices in Western QC and PEA panels.
What you should expect to see if it is working
Distinct, narrow target‑wise profiles in kDa with ladder‑based calibration error ≤ 10 percent over 15–250 kDa.
Linear response over 3–4 logs in single‑target tests after UMI collapse, with per‑bin CV ≤ 15 percent across technical replicates.
In the 24‑plex T‑cell panel, rank‑order agreement with Simple Western and phospho‑flow, with tighter replicate variance when using spike‑ins and total protein normalization.
Feasibility verdict
Technically feasible with careful chemistry and controls. The gating risk is background signal from oligo interactions and matrix inhibition of the polymerase. The second risk is biology specific, namely finding antibody pairs that both bind the same protein in lysate while tolerating the buffer you need. Both can be engineered down to acceptable levels with the mitigation stack below.
If your goal is accurate copy numbers for 10 to 20 proteins per sample in standard lysates, this can be achieved. For membrane proteins or harsh buffers, plan to use a bead capture step to remove inhibitors before PCR.
Key risks and how to retire them fast
Antibody binding after SDS Go/no‑go: binding at ≤0.01 percent SDS after cyclodextrin treatment must recover to at least 60–80 percent of native control for a test panel. If not, switch to a rapid buffer‑exchange or on‑bead SDS removal before binding.
Bin cross‑talk Quantify with a fluorescent ladder only. Require <3 percent signal in adjacent bins for the dominant band before moving on.
Barcode cross‑talk and index hopping Use unique dual indexes with error‑correcting antibody ID barcodes. Enforce strict adapter cleanup.
Insufficient effective resolution If dispersion broadens peaks across >3 bins at mid‑range, increase number of bins and shorten the separation window. Note that commercial chips deliver ∼10 percent resolution, which maps well to 64–128 bins over 15–250 kDa.
UMI collisions at high copy bins Use 14–16 nt UMIs and UMI‑aware deduplication tools.
Recommended MVP path
Prioritize Version A, plate‑based. It is the fastest credible build with standard gear.
Step 1. Build separation + fractionation stub
Microchip SDS run into a 96‑well plate. Pre‑aliquot bin barcodes across wells. Confirm bin‑to‑kDa calibration with a prestained ladder.
Acceptance: logistic fit r² ≥ 0.98, sizing error ≤ 10 percent for ladder bands.
Step 2. SDS neutralization check
Titrate β‑cyclodextrin in collected fractions and test one antibody–antigen pair known to be sensitive to SDS. Require ≥ 60 percent of native binding at ≤ 0.01 percent residual SDS.
Step 3. Single‑target end‑to‑end
One DNA‑tagged antibody, HeLa lysate spiked with recombinant target, 8‑point dilution, NGS readout with 16 nt UMIs. Expect linear counts over at least 3 logs with UMI collapse.
Step 4. Isoform resolution
PARP1 cleavage model. Show two non‑overlapping peaks at ∼116 and ∼89 kDa with <10 percent cross‑bin bleed.
Step 5. PTM specificity
ERK1/2 pT/pY with phosphatase control. Phospho signal should vanish in treated samples while total ERK remains. Use PEA for phospho to suppress off‑targets.
Step 6. 24‑plex T‑cell panel
Run your listed phospho panel. Benchmark fold‑changes against Simple Western and phospho‑flow. Expect rank‑order agreement and tighter CVs on DSI‑Seq if normalization is done with spike‑ins and per‑bin protein signal.
Quantitative design notes
Bins and segmentation: with a 2–3 minute separation window and 96–128 bins, you are only asking for ∼0.5–1.0 Hz effective binning. That is easy for a valve or droplet generator, and nanoliter per bin volumes are compatible with standard PCR chemistry. Reviews document CE‑to‑droplet interfaces.
Normalization hierarchy: 1) external spike‑in protein standards, 2) per‑bin total protein measurement via a dye read on the plate before PCR, 3) stable reference proteins.
Library design: error‑correcting 12–16 nt antibody IDs with Hamming distance ≥ 3, 14–16 nt UMIs, and unique dual indexes for samples.
Data analysis: UMI collapse, per‑bin background modeling, then peak calling on smoothed target‑wise bin profiles with FDR control. UMI‑aware tools will handle sequencing errors in UMIs.
Where to be extra hard on yourselves
SDS compatibility is the make‑or‑break. Literature is clear that SDS suppresses immunorecognition in solution. Beta‑cyclodextrin removal is credible, but you must prove quantitative recovery for several antibody classes and for phospho‑epitopes.
True size fidelity. Demand bin‑to‑kDa errors ≤ 10 percent across the ladder. Commercial microchip SDS assays hit this spec, so you should too.
Cross‑method agreement. In the 24‑plex test, require high concordance with Simple Western and targeted MS for several anchors. Targeted MS is the recognized Western alternative for quantitative work. Use it to certify your dynamic range and linearity.
Hard problems you must solve, with fixes
One protein should create one amplicon
Risk: proximity oligos extend or ligate without protein, or multivalent proteins give more than one product.
Fixes:
Use two oligos that are each locked in a short hairpin. A short connector strand opens both hairpins only when co‑localized on the same protein, then a high‑fidelity polymerase fills in to create the amplicon. Include 3′‑phosphate blocking until the connector is present.
For oligomers, design epitope pairs on a single monomer when possible. If not possible, report results per complex rather than per monomer and state that explicitly.
Antibody pair availability and behavior in lysate
Risk: lack of two non‑overlapping linear epitopes, steric clash, or loss of binding in detergent.
Fixes:
Start with clones validated for Western or IP, not only IF. Western and IP clones tolerate linear epitopes and exposure to detergents.
Perform epitope binning by BLI or SPR to choose non‑competing pairs separated by at least 5 to 10 nm on the unfolded or partially refolded protein model.
Where pairs are weak, replace one antibody with a nanobody or aptamer against a distinct linear epitope.
For phospho targets, prefer two antibodies that bind outside the modified residue, with one being modification specific. If this is impossible, move that target to capture‑and‑release.
Matrix inhibition of the proximity and PCR steps
Risk: detergents, salts, or protease inhibitors compromise extension and PCR.
Fixes:
Keep lysis buffer PCR compatible when possible. If you must use strong detergents, run capture‑and‑release: bead capture, wash to remove inhibitors, then release DNA reporter into clean PCR buffer.
Add a short post‑binding buffer exchange on magnetic beads for problematic targets.
Validate a polymerase mix that tolerates residual lysate at your intended dilution.
Background from carryover DNA or oligo self‑assembly
Risk: false positives dominate the low end and inflate the apparent copy number.
Fixes:
Pre‑clear the lysate with Benzonase, then inactivate it completely before proximity chemistry.
Use dUTP in all amplicons and add UDG before PCR to destroy any carryover products from previous runs.
Split pre‑PCR and post‑PCR spaces physically, with aerosol‑barrier tips and separate lab coats.
Sequence design: hairpin locks, high Hamming distance between barcodes, blocked ends, and no complementarity between any non‑partner oligos. Reject any panel where the reagent‑only control exceeds 0.2 positive partitions per 20,000 partitions.
Hook effect and nonlinearity at high abundance
Risk: signal per protein falls at high concentration because the two binders distribute on different molecules.
Fixes:
Titrate antibody concentrations into the zone where both epitopes are saturated for your expected protein range.
Always include a two‑point dilution series per sample for the abundant proteins. Accept a target only if the two dilutions agree within 10 percent after Poisson correction.
Multiplex cross‑talk
Risk: oligos from different pairs hybridize or amplify off target.
Fixes:
Build the panel by subpanels. Screen each new pair against the existing set for off‑target positives in a protein‑free background.
If a pair misbehaves, move it to the capture‑and‑release lane to isolate its chemistry.
Go or no‑go acceptance criteria
Adopt these hard thresholds before you believe a number.
Reagent‑only background
Less than 0.2 positives per 20,000 partitions per target channel.
Blank lysate after nuclease
Less than 0.5 positives per 20,000 partitions with universal primers but without proximity chemistry. If not, your nuclease or physical separation is insufficient.
Spike‑in recovery
Recovery 0.8 to 1.2 across a 100‑fold range for purified protein spiked into matrix. Failures trigger re‑titration of antibody concentrations and buffer cleanup.
Dilution linearity in real lysate
Two dilutions that differ by 2× must report a 1.8 to 2.2 ratio after Poisson correction.
Repeatability
Technical replicate CV less than 10 percent at 400 to 10,000 molecules per reaction.
Cross‑panel interference
Adding or removing other antibody pairs changes the measured copy number by less than 10 percent.
Bead‑gated proximityBind the capture antibody on beads, add the reporter antibody in solution, perform proximity extension on‑bead, then wash and elute the newly formed DNA into PCR buffer. This removes inhibitors and unreacted oligos and cuts background sharply. It reduces hands‑off simplicity but pays off for tough targets.
Connector‑mediated extension with dual locksEncode two short toeholds on the oligo arms and design a connector that bridges both with perfect Tm only when both arms are present. Include 3′ phosphates on arms and a 5′ phosphate on the connector. Use a strand‑displacing polymerase that starts only after ligation or nick‑translation. This reduces unspecific arm‑to‑arm pairing.
UDG carryover suppression and separate primer poolsUse dUTP in all products plus UDG in the PCR master mix. Maintain separate primer mixes for each subpanel to avoid building a universal contaminant that can amplify anywhere.
External protein counting standardInclude a recombinant protein with an artificial two‑epitope tag and its dedicated antibody pair in every reaction at a fixed copy number. This provides an internal process control for conversion efficiency and partition calling. Report all targets relative to this control and then convert back to absolute units with its known copy number.
Per‑target conversion efficiency calibrationMeasure epsilon for each target with a dilution series of purified protein in matrix. Store epsilon in software and apply a simple correction: estimated molecules equal measured molecules divided by epsilon. Update epsilon with each new antibody lot.
Panel design discipline
Keep amplicon length between 60 and 100 nt with matched Tm, design all probes against non‑overlapping internal sequences, and require Hamming distance of at least 3 for barcodes. Reject any oligo that creates primer dimers in silico against the entire panel.
PTM‑aware strategiesFor phosphorylation, when two antibodies cannot both bind, pair a pan antibody with a phospho‑specific binder and place the oligo on the pan arm only. Then use capture‑and‑release where phospho recognition triggers DNA release. This keeps the readout phospho dependent without needing both arms to bind.
Membrane protein handlingFor multi‑pass proteins, use mild detergents such as digitonin or DDM for extraction, then switch to bead capture with washes to remove detergents before the proximity step.
Failure modes you will see first, and how to debug in one day
Everything is positive
Likely contamination with prior amplicon. Replace all solutions, add UDG, and move pre‑ and post‑PCR physically apart. Confirm by running reagent‑only controls.
Nothing is positive, even with recombinant protein
Connector or polymerase step is not working, or one epitope is masked. Verify each arm can be extended alone with a synthetic splint, check antibody binding by classic sandwich ELISA, then return to proximity chemistry.
Strong signal in no‑protein control
Your arms are self‑complementary or the connector bridges unrelated arms. Redesign oligos with hairpins and toeholds that require co‑localization.
Nonlinear response across dilutions
You are in hook territory or conversion efficiency varies with protein concentration. Lower antibody concentration or split the sample. Re‑titrate to achieve flat efficiency across the intended range.
Statistics you should report with every result
Molecules per reaction with 95 percent confidence intervals based on binomial uncertainty of positives, converted via the standard Poisson occupancy model.
Dilution agreement score for each target.
Conversion efficiency epsilon and its uncertainty if you apply the correction.
Background positives per 20,000 partitions for the run.
Practical build choices I recommend
Start plate‑based. Use a microchip SDS instrument or capillary interface to a 96‑well plate, pre‑aliquot bin barcodes, do all chemistry in wells. This gets you data with minimal custom microfluidics. Published interfaces from electrophoresis to droplets or plates give you patterns to copy.
PEA for phospho targets, single‑epitope capture for abundant structural targets. PEA is proven to reduce off‑target noise in multiplex settings.
SDS removal: add β‑cyclodextrin and a brief dilution step upon fraction entry, then bind. Validate residual SDS by a colorimetric assay or mass‑balance against standards.
Controls: isotypes, no‑antibody bins, phosphatase controls for phospho, and known‑ratio recombinant mixes to certify linearity. This mirrors best practices in Western QC and PEA panels.
What you should expect to see if it is working
Distinct, narrow target‑wise profiles in kDa with ladder‑based calibration error ≤ 10 percent over 15–250 kDa.
Linear response over 3–4 logs in single‑target tests after UMI collapse, with per‑bin CV ≤ 15 percent across technical replicates.
In the 24‑plex T‑cell panel, rank‑order agreement with Simple Western and phospho‑flow, with tighter replicate variance when using spike‑ins and total protein normalization.
Below is a focused, execution‑ready plan to retire the highest risks in DSI‑Seq fast, with concrete tests, acceptance gates, and fixes. It keeps the assay in the IP‑lean, single‑tag digital immunoPCR architecture you asked for.
This plan gives you fast, quantitative readouts for each major risk and a clear go or no‑go at every stage. If you want, I can turn the SDS titration and cross‑talk assays into two mini SOPs with prefilled data sheets so your team can run them tomorrow and record pass or fail unambiguously.
2) Antibody binding after SDS
Goal
Prove that antibody binding in collected fractions is acceptable after SDS scavenging, or switch to a short buffer‑exchange on beads without sacrificing throughput.
Fast experiment (same day)
Plate setup: Prepare eight sets of collection buffer with 0, 5, 10, 15, 20, 30, 40, 60 mM β‑cyclodextrin, all with 2 mg mL⁻¹ BSA and 0.05 percent Tween‑20.
Separation: Run SDS‑CE on a test lysate plus ladder, collect 24 bins into each cyclodextrin condition.
Residual SDS readout: In a spare row, measure SDS per well by MBAS assay or equivalent colorimetric surfactant assay. Acceptance: residual SDS ≤0.01 percent at or below 20 mM cyclodextrin for the mid‑run bins.
Binding panel: In each condition, test six antibodies that represent your real use: two phospho‑specific, two linear epitope, one conformational, one membrane‑proximal. Perform standard capture on beads followed by detector binding and single‑tag release. Quantify tags by dPCR to keep turnaround short.
Acceptance gate: At least 5 of 6 antibodies recover 60–80 percent of native binding at ≤0.01 percent residual SDS.
If fail
Bead‑based buffer exchange per bin: after fraction collection, incubate with capture beads 15 minutes, pull down, wash 3 times in PBS‑Tween, then add detector. This removes SDS physically before detector binding.
Detergent‑removal spin columns: 96‑well cartridges to remove SDS from fractions in 2–3 minutes.
Collection buffer upgrade: raise β‑cyclodextrin to 30–40 mM and add 0.1 percent Triton X‑100 to reduce protein sticking while scavenging SDS; re‑test.
Lock‑in test
When a recipe passes, re‑run the six‑antibody panel on three different lysate types and two phospho states. Keep the passing cyclodextrin concentration as a default.
3) Bin cross‑talk
Measurement
Run a fluorescent ladder. Collect 48 bins at 15 seconds each. Measure per‑bin fluorescence on a plate reader.
For the tallest band, compute:
central bin area A₀,
adjacent bins A₋₁ and A₊₁,
cross‑talk percent = 100 × (A₋₁ + A₊₁) / A₀.
Acceptance
Less than 3 percent combined in the two adjacent bins.
Rapid fixes
Waste gap: between bins, move outlet to a waste well for 2 seconds to dump the tail.
Shorten dead volume: minimize outlet tubing length, or bring the capillary tip to touch each well’s meniscus.
Increase segmentation rate: reduce bin width to 10 seconds; keep total window length by adding bins.
Tighten timing: verify stage moves with a stopwatch; jitter should be less than ±0.1 second.
Re‑measure after each change until the gate is met.
4) Barcode cross‑talk and index hopping
Library design
Antibody ID tags: 12–16 nt, Hamming distance at least 3.
Unique dual indexes for samples and for bin indexing. Never reuse an index pair within a run.
Controls to include in every NGS pool
Dark indexes: 8 index combinations that are not assigned to any bin. They report pure hopping.
No‑template bins: 4 wells processed without tags. They report adapter carryover and misassignment.
Acceptance
Index hopping: less than 0.2 percent for each dark index.
Off‑panel tag reads: less than 0.2 percent of total assigned reads.
Fixes if above thresholds
Double‑SPRI cleanup: 0.8x then 1.2x to remove free adapters.
Exonuclease I treatment after indexing PCR to remove leftover primers.
Reduce cluster density by 10–15 percent.
Increase bin‑specific index diversity to reduce miscalls by the demultiplexer.
5) Effective resolution
What to measure
From ladder bins, fit a logistic or quadratic time‑to‑logMW mapping. Record r² and per‑band sizing error.
For a mid‑range band near 50–70 kDa, compute the number of bins across the FWHM of its peak.
Acceptance
r² at least 0.98; sizing error at most 10 percent per ladder band.
FWHM spans no more than 3 bins.
Levers to improve
Increase field strength modestly and shorten the separation window to reduce diffusion.
Raise bin count to 64–128 while keeping the total window 10–15 minutes.
Refresh or slightly increase sieving polymer concentration.
Keep temperature stable; a 2 C drift will degrade mapping.
6) UMI collisions at high‑copy bins
UMI space is 4^L. Collisions rise with molecule count per bin.
Expected duplicate fraction (approximate):
With 14 nt UMIs (268,435,456 codes):
100,000 molecules: about 0.037 percent duplicates
500,000 molecules: about 0.186 percent
1,000,000 molecules: about 0.373 percent
With 16 nt UMIs (4,294,967,296 codes):
100,000 molecules: about 0.0023 percent
500,000 molecules: about 0.0116 percent
1,000,000 molecules: about 0.023 percent
Policy:
Use 14 nt UMIs for expected per‑bin molecule counts under 200,000.
Use 16 nt UMIs for anything higher.
Always deduplicate with UMI‑aware tools that correct one sequencing error in the UMI and collapse exact duplicates.
7) Recommended MVP path, expanded
Step 1. Separation plus fractionation stub
Run microchip SDS into a 96‑well plate with 48 bins at 15 seconds each.
Pre‑aliquot bin index adapters in each well.
Calibrate bin‑to‑kDa with a prestained ladder.
Acceptance: logistic fit r² at least 0.98; ladder sizing error at most 10 percent.
Step 2. SDS neutralization check
Titrate β‑cyclodextrin as in Section 2 and run the six‑antibody test.
Acceptance: at least 60 percent of native binding at ≤0.01 percent residual SDS.
Step 3. Single‑target end‑to‑end
One target, HeLa lysate spiked with purified protein, 8‑point dilution, NGS readout, 16 nt UMIs.
Expect: linear counts over at least 3 logs after UMI collapse; reagent‑only background under 0.2 percent of total reads.
Step 4. Isoform resolution
PARP1 cleavage model. Two non‑overlapping peaks at about 116 and about 89 kDa.
Acceptance: less than 10 percent cross‑bin bleed between the two peaks, peak FWHM within 3 bins.
Step 5. PTM specificity
ERK1/2 pT/pY with a phosphatase control.
IP‑safe detection: capture with a pan ERK antibody on beads, detect with a phospho‑specific single‑tag detector. No proximity ligation or extension is used.
Acceptance: phospho signal falls to background in phosphatase‑treated samples while total ERK profile remains.
Step 6. 24‑plex T‑cell panel
Run the 24‑plex DSI‑Seq panel by NGS.
Benchmark fold‑changes against capillary Western and phospho‑flow on the same samples.
Expect: rank‑order agreement across targets. With spike‑ins and per‑bin total‑protein normalization, DSI‑Seq should yield tighter CVs than Western.
8) Quantitative design notes to enforce
Bins and segmentation: for a 12–15 minute window and 96–128 bins you are at 0.5–1.0 Hz. This is easy for a plate stage or a droplet generator. Keep outlet dead volume minimal.
Normalization hierarchy: prioritize (1) external protein spike‑ins per sample, (2) per‑bin total protein dye read on the plate before PCR, (3) a stable reference protein.
Library design: 12–16 nt antibody IDs with Hamming distance at least 3; 16 nt UMIs for high bins; unique dual indexes for samples and for bins; strict adapter cleanup.
Data analysis: UMI collapse with single‑error correction, per‑bin background modeling from reagent‑only wells, peak calling on smoothed target profiles with FDR control across bins.
9) Execution timeline
Day 1 morning
Calibrate stage timing, verify outlet alignment, prepare cyclodextrin plates.
Conjugate one detector and QC 1:1 stoichiometry.
Day 1 afternoon
Run ladder and lysate fractions across cyclodextrin titration.
Measure residual SDS and run the six‑antibody binding panel.
If acceptable, lock cyclodextrin. If not, add bead‑based buffer exchange and repeat a subset.
Day 2
Single‑target spike‑in series, NGS library with 16 nt UMI, run on a desktop sequencer.
Process reads, check linearity and background.
Day 3–4
PARP1 cleavage model and ERK phospho specificity.
Re‑measure cross‑talk and effective resolution after any mechanical tweaks.
Day 5–7
Assemble 24‑plex and run two biological samples with orthogonal comparators.
10) Extra scrutiny for SDS compatibility
Be strict here. SDS suppression of immunorecognition is well known. Your acceptance is recovery of 60–80 percent of native binding at or below 0.01 percent residual SDS across several antibody classes. If you cannot hit that, standardize on bead capture in each bin followed by washing before detector binding. It adds 10–15 minutes but removes the single biggest source of failure.
11) Acceptance checklist to sign per run
Reagent‑only background under 0.2 percent of reads or under 0.2 positives per 20,000 partitions (dPCR).
Index hopping under 0.2 percent on dark indexes.
Ladder r² at least 0.98; per‑band error at most 10 percent.
Peak FWHM within 3 bins at mid‑range.
Cross‑bin bleed under 3 percent for dominant band.
UMI collision policy satisfied given observed molecule counts.
Spike‑in recovery 0.8–1.2 across a 100‑fold range.
Two‑point dilution linearity 1.8–2.2× for a 2× input change.
12) When to pivot
If SDS compatibility fails after bead exchange, pivot to native separation for targets that tolerate it and keep SDS only for targets that require denaturation.
If bin cross‑talk persists after waste gaps and timing fixes, switch to droplet segmentation at 1–2 Hz with an in‑line waste gap every 1–2 droplets.
If index hopping remains above 0.2 percent, use a platform with patterned‑flow‑cell mitigations for hopping or reduce on‑instrument carryover by splitting the run.
This plan gives you fast, quantitative readouts for each major risk and a clear go or no‑go at every stage. If you want, I can turn the SDS titration and cross‑talk assays into two mini SOPs with prefilled data sheets so your team can run them tomorrow and record pass or fail unambiguously.
2) Antibody binding after SDS
Goal
Prove that antibody binding in collected fractions is acceptable after SDS scavenging, or switch to a short buffer‑exchange on beads without sacrificing throughput.
Fast experiment (same day)
Plate setup: Prepare eight sets of collection buffer with 0, 5, 10, 15, 20, 30, 40, 60 mM β‑cyclodextrin, all with 2 mg mL⁻¹ BSA and 0.05 percent Tween‑20.
Separation: Run SDS‑CE on a test lysate plus ladder, collect 24 bins into each cyclodextrin condition.
Residual SDS readout: In a spare row, measure SDS per well by MBAS assay or equivalent colorimetric surfactant assay. Acceptance: residual SDS ≤0.01 percent at or below 20 mM cyclodextrin for the mid‑run bins.
Binding panel: In each condition, test six antibodies that represent your real use: two phospho‑specific, two linear epitope, one conformational, one membrane‑proximal. Perform standard capture on beads followed by detector binding and single‑tag release. Quantify tags by dPCR to keep turnaround short.
Acceptance gate: At least 5 of 6 antibodies recover 60–80 percent of native binding at ≤0.01 percent residual SDS.
If fail
Bead‑based buffer exchange per bin: after fraction collection, incubate with capture beads 15 minutes, pull down, wash 3 times in PBS‑Tween, then add detector. This removes SDS physically before detector binding.
Detergent‑removal spin columns: 96‑well cartridges to remove SDS from fractions in 2–3 minutes.
Collection buffer upgrade: raise β‑cyclodextrin to 30–40 mM and add 0.1 percent Triton X‑100 to reduce protein sticking while scavenging SDS; re‑test.
Lock‑in test
When a recipe passes, re‑run the six‑antibody panel on three different lysate types and two phospho states. Keep the passing cyclodextrin concentration as a default.
3) Bin cross‑talk
Measurement
Run a fluorescent ladder. Collect 48 bins at 15 seconds each. Measure per‑bin fluorescence on a plate reader.
For the tallest band, compute:
central bin area A₀,
adjacent bins A₋₁ and A₊₁,
cross‑talk percent = 100 × (A₋₁ + A₊₁) / A₀.
Acceptance
Less than 3 percent combined in the two adjacent bins.
Rapid fixes
Waste gap: between bins, move outlet to a waste well for 2 seconds to dump the tail.
Shorten dead volume: minimize outlet tubing length, or bring the capillary tip to touch each well’s meniscus.
Increase segmentation rate: reduce bin width to 10 seconds; keep total window length by adding bins.
Tighten timing: verify stage moves with a stopwatch; jitter should be less than ±0.1 second.
Re‑measure after each change until the gate is met.
4) Barcode cross‑talk and index hopping
Library design
Antibody ID tags: 12–16 nt, Hamming distance at least 3.
Unique dual indexes for samples and for bin indexing. Never reuse an index pair within a run.
Controls to include in every NGS pool
Dark indexes: 8 index combinations that are not assigned to any bin. They report pure hopping.
No‑template bins: 4 wells processed without tags. They report adapter carryover and misassignment.
Acceptance
Index hopping: less than 0.2 percent for each dark index.
Off‑panel tag reads: less than 0.2 percent of total assigned reads.
Fixes if above thresholds
Double‑SPRI cleanup: 0.8x then 1.2x to remove free adapters.
Exonuclease I treatment after indexing PCR to remove leftover primers.
Reduce cluster density by 10–15 percent.
Increase bin‑specific index diversity to reduce miscalls by the demultiplexer.
5) Effective resolution
What to measure
From ladder bins, fit a logistic or quadratic time‑to‑logMW mapping. Record r² and per‑band sizing error.
For a mid‑range band near 50–70 kDa, compute the number of bins across the FWHM of its peak.
Acceptance
r² at least 0.98; sizing error at most 10 percent per ladder band.
FWHM spans no more than 3 bins.
Levers to improve
Increase field strength modestly and shorten the separation window to reduce diffusion.
Raise bin count to 64–128 while keeping the total window 10–15 minutes.
Refresh or slightly increase sieving polymer concentration.
Keep temperature stable; a 2 C drift will degrade mapping.
6) UMI collisions at high‑copy bins
UMI space is 4^L. Collisions rise with molecule count per bin.
Expected duplicate fraction (approximate):
With 14 nt UMIs (268,435,456 codes):
100,000 molecules: about 0.037 percent duplicates
500,000 molecules: about 0.186 percent
1,000,000 molecules: about 0.373 percent
With 16 nt UMIs (4,294,967,296 codes):
100,000 molecules: about 0.0023 percent
500,000 molecules: about 0.0116 percent
1,000,000 molecules: about 0.023 percent
Policy:
Use 14 nt UMIs for expected per‑bin molecule counts under 200,000.
Use 16 nt UMIs for anything higher.
Always deduplicate with UMI‑aware tools that correct one sequencing error in the UMI and collapse exact duplicates.
7) Recommended MVP path, expanded
Step 1. Separation plus fractionation stub
Run microchip SDS into a 96‑well plate with 48 bins at 15 seconds each.
Pre‑aliquot bin index adapters in each well.
Calibrate bin‑to‑kDa with a prestained ladder.
Acceptance: logistic fit r² at least 0.98; ladder sizing error at most 10 percent.
Step 2. SDS neutralization check
Titrate β‑cyclodextrin as in Section 2 and run the six‑antibody test.
Acceptance: at least 60 percent of native binding at ≤0.01 percent residual SDS.
Step 3. Single‑target end‑to‑end
One target, HeLa lysate spiked with purified protein, 8‑point dilution, NGS readout, 16 nt UMIs.
Expect: linear counts over at least 3 logs after UMI collapse; reagent‑only background under 0.2 percent of total reads.
Step 4. Isoform resolution
PARP1 cleavage model. Two non‑overlapping peaks at about 116 and about 89 kDa.
Acceptance: less than 10 percent cross‑bin bleed between the two peaks, peak FWHM within 3 bins.
Step 5. PTM specificity
ERK1/2 pT/pY with a phosphatase control.
IP‑safe detection: capture with a pan ERK antibody on beads, detect with a phospho‑specific single‑tag detector. No proximity ligation or extension is used.
Acceptance: phospho signal falls to background in phosphatase‑treated samples while total ERK profile remains.
Step 6. 24‑plex T‑cell panel
Run the 24‑plex DSI‑Seq panel by NGS.
Benchmark fold‑changes against capillary Western and phospho‑flow on the same samples.
Expect: rank‑order agreement across targets. With spike‑ins and per‑bin total‑protein normalization, DSI‑Seq should yield tighter CVs than Western.
8) Quantitative design notes to enforce
Bins and segmentation: for a 12–15 minute window and 96–128 bins you are at 0.5–1.0 Hz. This is easy for a plate stage or a droplet generator. Keep outlet dead volume minimal.
Normalization hierarchy: prioritize (1) external protein spike‑ins per sample, (2) per‑bin total protein dye read on the plate before PCR, (3) a stable reference protein.
Library design: 12–16 nt antibody IDs with Hamming distance at least 3; 16 nt UMIs for high bins; unique dual indexes for samples and for bins; strict adapter cleanup.
Data analysis: UMI collapse with single‑error correction, per‑bin background modeling from reagent‑only wells, peak calling on smoothed target profiles with FDR control across bins.
9) Execution timeline
Day 1 morning
Calibrate stage timing, verify outlet alignment, prepare cyclodextrin plates.
Conjugate one detector and QC 1:1 stoichiometry.
Day 1 afternoon
Run ladder and lysate fractions across cyclodextrin titration.
Measure residual SDS and run the six‑antibody binding panel.
If acceptable, lock cyclodextrin. If not, add bead‑based buffer exchange and repeat a subset.
Day 2
Single‑target spike‑in series, NGS library with 16 nt UMI, run on a desktop sequencer.
Process reads, check linearity and background.
Day 3–4
PARP1 cleavage model and ERK phospho specificity.
Re‑measure cross‑talk and effective resolution after any mechanical tweaks.
Day 5–7
Assemble 24‑plex and run two biological samples with orthogonal comparators.
10) Extra scrutiny for SDS compatibility
Be strict here. SDS suppression of immunorecognition is well known. Your acceptance is recovery of 60–80 percent of native binding at or below 0.01 percent residual SDS across several antibody classes. If you cannot hit that, standardize on bead capture in each bin followed by washing before detector binding. It adds 10–15 minutes but removes the single biggest source of failure.
11) Acceptance checklist to sign per run
Reagent‑only background under 0.2 percent of reads or under 0.2 positives per 20,000 partitions (dPCR).
Index hopping under 0.2 percent on dark indexes.
Ladder r² at least 0.98; per‑band error at most 10 percent.
Peak FWHM within 3 bins at mid‑range.
Cross‑bin bleed under 3 percent for dominant band.
UMI collision policy satisfied given observed molecule counts.
Spike‑in recovery 0.8–1.2 across a 100‑fold range.
Two‑point dilution linearity 1.8–2.2× for a 2× input change.
12) When to pivot
If SDS compatibility fails after bead exchange, pivot to native separation for targets that tolerate it and keep SDS only for targets that require denaturation.
If bin cross‑talk persists after waste gaps and timing fixes, switch to droplet segmentation at 1–2 Hz with an in‑line waste gap every 1–2 droplets.
If index hopping remains above 0.2 percent, use a platform with patterned‑flow‑cell mitigations for hopping or reduce on‑instrument carryover by splitting the run.
I am also adding an organized lab‑ready protocol for Digital Size‑Indexed ImmunoSequencing (DSI‑Seq) here.
This is written to let you prototype, optimize, and validate the method on benchtop equipment. The protocol implements the IP‑lean single‑tag digital immunoPCR style readout inside each size bin, and it avoids proximity ligation or proximity extension chemistry. It supports two readouts:
NGS counting for high multiplex and practical bin indexing.
dPCR counting for small target panels or early debugging.
The workflow is plate‑based, since that is the fastest and most accessible path to working data.
0. Scope and outcome
Goal. Resolve proteins by apparent size into many narrow fractions, then quantify specific targets in each fraction by converting each captured protein into one DNA tag and counting tags. Output is a matrix of counts: targets by size bins, calibrated to kDa using a co‑run ladder.
Typical first build. 48 bins across a 12 minute separation window, 12 to 24 targets by NGS readout, input 1 to 10 micrograms total protein per lane, results in one day after library prep.
1. Safety and handling
Acrylamide and SDS are irritants. Handle with gloves, lab coat, and eye protection. Dispose of acrylamide waste as hazardous.
UV photocleavage variants require eye and skin protection. Shield the UV source.
Nucleases can aerosolize. Open tubes slowly and decontaminate surfaces.
Segregate pre‑PCR and post‑PCR spaces. Use UDG carryover control.
2. Materials and reagents
2.1 Buffers and solutions
Lysis buffer, PCR compatible50 mM HEPES pH 7.5, 150 mM NaCl, 1 percent NP‑40 or 0.5 percent Triton X‑100, 10 percent glycerol, protease inhibitors, phosphatase inhibitors if needed.
Optional SDS denaturing load buffer for chip1x TGS or TBE, 0.1 to 0.5 percent SDS, 10 mM DTT, 10 percent glycerol. Heat 70 to 90 C for 5 minutes if full denaturation is required.
Replaceable sieving polymer for SDS‑CECommercial SDS‑CE polymer or 2.5 to 4.0 percent linear polyacrylamide in 1x TBE with 0.1 percent SDS.
Collection buffer per well20 mM HEPES pH 7.5, 150 mM NaCl, 0.05 percent Tween‑20, 10 mM methyl‑beta‑cyclodextrin, 2 mg mL‑1 BSA. The cyclodextrin scavenges SDS on contact.
Bead capture bufferPBS, 0.05 percent Tween‑20, 1 mg mL‑1 BSA.
Wash bufferPBS, 0.05 percent Tween‑20.
Cleavage bufferDepends on linker. For disulfide linkers: PBS with 10 to 50 mM TCEP. For o‑nitrobenzyl photocleavage: PBS.
UDG mix for carryover controldNTPs with dUTP substitution, thermolabile UDG.
Benzonase mix25 to 50 U mL‑1 Benzonase with 2 mM MgCl2. Stop with 10 mM EDTA or heat inactivation per supplier.
2.2 Controls and standards
Protein ladder 10 to 250 kDa, prestained.
Spike‑in protein standards for at least two targets, quantified concentration.
Negative control lysate with phosphatase treatment for PTM assays.
2.3 Antibodies and DNA tags
Capture antibodies coupled to magnetic beadsUse Protein G beads with oriented coupling or covalent coupling kits. Aim for 1 to 5 micrograms antibody per mg beads. Prepare one capture bead type per target.
Detector binders that are monovalent with one DNA tag eachPrefer Fab fragments or nanobodies engineered with a single cysteine. Conjugate one DNA tag per binder by maleimide‑thiol chemistry. Confirm 1:1 stoichiometry by intact mass.
DNA tag designLength 60 to 100 nt.Layout: 5’ universal priming site A, 8 to 12 nt UMI, 12 to 16 nt target ID, 5’ half of an Illumina adapter if using NGS, 3’ blocking group if needed.The tag is attached to the detector via a generic, cleavable linker. No two‑probe hybridization is used.
Per‑bin index adapters for NGSShort double‑stranded adapters with bin‑specific index sequences and universal priming sites. These are ligated or appended in PCR to encode bin identity.
TaqMan probes for dPCROne probe per target tag. For dPCR you cannot multiplex many bins in one tube, so plan per‑bin reactions only for 1 to 4 targets.
2.4 Equipment
Microchip or capillary SDS electrophoresis instrument with replaceable polymer.
Simple fraction collector setupA motorized XY stage for a 96‑well plate under the capillary outlet. Alternatively a commercial fraction collector with programmable step time.
Magnetic racks for 96‑well plates.
Thermocycler and a bench NGS library prep kit.
dPCR instrument if using the dPCR readout.
Plate shaker, microcentrifuge, fluorometer or qPCR for library QC.
Optional UV source for photocleavage, with shielding.
3. Panel design and conjugation
Choose targets. Begin with 12 to 24 proteins. Favor clones validated for Western or IP. Include one reference housekeeper.
Epitope check. Select capture and detector epitopes that do not compete. For oligomers or large complexes, expect a calibration factor as noted later.
Conjugate detectors.
Reduce Fab or nanobody to expose the engineered cysteine.
React with maleimide‑tagged DNA at 1.5 to 2.0 molar excess.
Purify by size exclusion or affinity to remove free DNA.
Confirm 1:1 by intact mass or denaturing CE.
Bead coupling. Couple capture antibody to magnetic beads. Block with BSA. Store in PBS with 0.02 percent sodium azide at 4 C.
Document the per‑target conversion factor ρ. If two detectors can bind one captured protein or the protein is oligomeric, measure ρ using purified protein and record it in the target sheet.
4. Sample preparation
Harvest cells or tissue. Keep cold.
Lyse in PCR‑compatible lysis buffer. For maximum size resolution across proteoforms you may choose an SDS denaturing load immediately before separation.
Clarify by spinning at 14,000 g for 10 minutes.
Protein quant by BCA.
Nuclease treatment for protein‑only readout. Add Benzonase mix to lysate, incubate 15 minutes at room temperature. Stop with EDTA or heat inactivate. This ensures no endogenous DNA or RNA is counted.
Spike‑in controls. Add defined copies of two recombinant controls.
Add ladder for a dedicated ladder run. Prepare a separate tube with the prestained ladder in the same matrix for calibration runs.
5. Microchip SDS separation
You can run native or denaturing. For size indexing you will typically run SDS.
Prime the chip with replaceable sieving polymer per instrument instructions.
Equilibrate with 1x running buffer containing 0.1 percent SDS.
Load sample. If using SDS, mix lysate with SDS load buffer to 0.1 to 0.5 percent SDS and heat 70 to 90 C for 5 minutes.
Injection. Electrokinetic inject for 3 to 5 seconds at reduced field.
Separation. Field strength 200 to 300 V cm‑1, total separation window 10 to 15 minutes. Record the exact time zero when injection ends.
Run a separate ladder trace with the same separation program and collect fractions the same way as the sample run.
6. Plate‑based fractionation into size bins
Prepare a 96‑well plate. Dispense 10 to 15 microliters of collection buffer into the wells you will use.
Define bins. For a 12 minute separation choose 48 bins at 15 seconds per bin. That uses half a 96‑well plate.
Position the capillary outlet above the first well. Use a droplet‑friendly tip or align to touch the meniscus.
Collect. Start separation and move the plate on a schedule so that each bin collects exactly 15 seconds of eluent. A simple script on the stage controller is enough.
Finish. Seal the plate. The cyclodextrin in the collection buffer reduces free SDS below 0.02 percent in seconds and helps renaturation of linear epitopes.
Optional fraction concentration. If fractions are very dilute, you can add 1 mg of dry Bio‑Beads SM‑2 per well for 5 minutes to further scavenge SDS, then remove beads with a magnetic wand or by decanting. Validate that your targets are not lost.
7. Per‑bin immunoassay with single‑tag detectors
This yields one DNA tag per captured protein complex in each bin. No proximity ligation or extension is used.
For each bin well:
Add capture beads
Add 10 to 20 microliters of bead suspension with the capture antibody for target i, or use a bead mix containing distinct bead codes if you run targets in parallel within a bin.
Incubate 30 minutes at room temperature with gentle shaking.
Magnet and wash
Pull beads down. Remove supernatant.
Wash 3 times with 100 microliters wash buffer.
Detector binding
Add the monovalent detector conjugate carrying a single DNA tag, at 1 to 5 nM in bead capture buffer.
Incubate 30 minutes.
Wash 3 times.
Release DNA tags
Add 20 microliters cleavage buffer.
For disulfide linkers, incubate with 10 to 50 mM TCEP for 10 minutes.
For photocleavage, illuminate with the specified UV for the vendor’s o‑nitrobenzyl linker. Keep temperature controlled.
Collect the supernatant which contains the released DNA tags.
UDG carryover control
If you use dUTP in tags, add thermolabile UDG and incubate per supplier before amplification to remove any carryover products. The released tags do not contain prior amplicons.
Pooling strategy
For NGS readout keep each bin separate through the next indexing step.
For dPCR readout proceed per bin per target. dPCR is best limited to 1 to 4 targets across a small number of bins during development.
Notes
If you prefer, perform steps 1 to 3 in bulk per bin with a mixed bead cocktail for all targets. This speeds handling. The detector step remains one DNA per bound protein because each detector is monovalent.
If an antigen accepts two detectors per captured protein, apply the per‑target factor ρ during analysis.
8. Readout A. NGS library and sequencing
NGS is the practical route for many bins and many targets because you can encode bin identity by indexing.
Per‑bin index addition
Set up a PCR using primers that add a unique bin index i5 or i7 to each bin. 10 to 14 cycles are typical since the tag pool can be low copy.
Use a common reverse primer so that only the index changes across bins.
Target identity in the tag
The target ID is already encoded in the tag sequence. You do not add any probe hybridization step between oligos on two antibodies. You simply amplify what you cleaved.
Pool and clean up
Pool equal volumes or equal mass from all bin PCRs for a sample.
Clean up with SPRI beads at 1.2x ratio.
QC and run
Measure library size and concentration. Expect 150 to 300 bp.
Sequence paired‑end or single‑end, 50 to 100 cycles is usually enough.
Depth target per sample 1 to 3 million reads for a 24 target by 48 bin design.
Demultiplex
Demultiplex by sample and by bin index.
Parse reads to tag IDs and UMIs. Collapse UMIs to unique molecules.
9. Readout B. dPCR per bin for 1 to 4 targets
Use this for early method checkout or when you only need a few targets.
Set up dPCR reactions
For each bin and each target, set up a reaction with the target‑specific TaqMan probe.
Partition and run endpoint PCR.
Count
Record positive partitions k and total partitions N.
Compute lambda per bin as λ = −ln(1 − k/N).
Molecules per bin equals λ times N.
Scale
Apply the per‑target calibration factor ρ if relevant.
Continue to the size calibration step below.
10. Size calibration using the ladder
You will convert bin index to apparent kDa.
Collect a ladder run with identical fractionation
Run the prestained ladder alone with the same separation and fraction schedule used for the sample. Collect into a plate with the same 48 bins.
Quantify ladder protein abundance in each bin by simple absorbance at 595 nm if dye permits, or by a quick fluorometric protein assay across wells.
Identify peaks
Ladder proteins will appear as peaks across bins. Fit peak centers to their known molecular weights.
Fit the mapping
A robust model is: log10(MW) = a + b·t + c·t^2 where t is bin center time in minutes.
Fit a, b, c by least squares using the ladder peaks.
For each bin j in the sample, compute its center time and then its apparent kDa by inverting the relationship.
Uncertainty
Compute 95 percent confidence intervals on MW per bin from the residuals of the fit.
11. Data processing and outputs
Counts table
For NGS, produce a table of unique molecule counts per target per bin after UMI collapse.
For dPCR, produce molecules per bin from Poisson correction.
Normalize
Divide by the external spike‑in recovery to correct for run‑to‑run variation.
Optionally normalize per bin by a total protein signal if measured.
Proteoform calling
For each target, smooth the binned profile with a small window.
Identify local maxima as proteoform peaks.
Report peak bin, apparent kDa, and integrated counts under each peak.
Final report
Matrix: targets × bins with counts.
Proteoform summary table with apparent kDa, counts, fraction of total per target.
QC panel with ladder fit, spike‑in recovery, background levels, replicate CVs.
12. Acceptance criteria and QC
Adopt these hard thresholds before you trust a run.
Reagent‑only background. After full library prep with no protein, fewer than 0.2 percent of reads assign to valid tag IDs or fewer than 0.2 positives per 20,000 partitions for dPCR.
Blank lysate after nuclease. With universal primers but no released tags, signal must be at background.
Spike‑in recovery. 0.8 to 1.2 across a 100‑fold range.
Dilution linearity. Two dilutions at 2x load must yield 1.8 to 2.2x counts per bin.
Ladder fit residual. Standard error of log10(MW) fit less than 0.03.
Replicates. Technical replicate CV less than 10 percent in bins with at least 400 molecules.
13. Validation plan
Run in this sequence.
Separation and fractionation stub
Ladder only. Build the bin to kDa mapping and verify timing precision over three runs.
Single‑target end‑to‑end
One target in a control lysate. Spike a dilution series of the purified protein. Validate linearity and the expected single peak at the known kDa.
Isoform resolution
Use a biology that creates a known cleavage, for example PARP1 cleavage in apoptosis. Confirm two peaks at expected apparent sizes.
PTM specificity
Use a phospho target with a modification specific detector. Compare stimulated vs phosphatase treated. The modified target should vanish in treated samples without a shift in the pan profile.
Multiplex cross‑talk
Build the panel in subpanels of 6 to 8 targets. Check background in no‑antigen wells and confirm no off‑target rises when subpanels are combined.
Reproducibility and lot stability
Ten technical replicates. Track per‑bin per‑target CV.
New detector lots get a check of the conversion factor ρ and a small ladder redo.
14. Troubleshooting
Weak or no signal in all binsCheck detector conjugation efficiency and cleavage step. Confirm 1:1 stoichiometry. Verify that the UDG step is not destroying tags.
High background in no‑protein controlsSuspect amplicon carryover or free DNA tag contamination. Increase physical separation of spaces, add UDG, and re‑purify detectors to remove free tag.
Flat size profile without peaksFraction timing off or excessive diffusion. Shorten bin width to 10 seconds or increase field strength. Confirm collection buffer is dispensed before the run.
Shifted ladder mapping between runsTemperature or polymer batch differences. Always run a ladder with each batch and refit a, b, c. Apply mapping per run.
Loss of antibody binding due to SDSIncrease cyclodextrin concentration in collection buffer to 20 mM. Add a short bead‑based buffer exchange before detector binding. Use clones validated for linear epitopes.
15. Options and upgrades
Droplet fractionationReplace the plate with a T‑junction droplet maker that encapsulates the outlet stream at a fixed rate. A side stream can inject a droplet index oligo that is appended later by standard ligation. This increases bin count to 96 to 128 at the same run time.
Bin reduction mergeFor abundant targets, merge adjacent bins in software to increase counts per bin and improve precision.
Alternative SDS scavengersPotassium chloride can precipitate SDS as potassium dodecyl sulfate. Use with care due to protein precipitation. Validate recovery on spike‑ins.
16. IP and compliance posture
One DNA tag per detector binder.
No proximity ligation or proximity extension between two nucleic acid‑bearing probes.
No single‑molecule enzyme arrays.
Bin identity is appended during library prep by standard indexing, or dPCR is run per bin per target without any probe‑probe interaction.
Include UDG carryover suppression and commodity linkers only.
Record these choices in your design history file from day one.
17. Starter T cell panel
Begin with 12 to 18 targets that are Western or IP validated and bind linear epitopes. For example: CD3ζ, ZAP70, LAT, SLP76, PLCG1, ERK1, ERK2, AKT, mTOR, 4EBP1, NF‑κB p65, PARP1, and a reference such as beta‑actin. Add phospho‑specific detectors as a separate subpanel and always run a phosphatase control.
18. Example day plan
Day 1 morning
Conjugate 2 to 4 detectors and QC one by mass.
Couple 2 to 4 capture antibodies to beads and block.
Day 1 afternoon
Prepare lysates, nuclease treat, spike controls.
Run ladder and sample separations with fractionation to 48 bins.
Day 1 evening
Perform per‑bin bead capture, detector binding, cleavage.
Start per‑bin index PCR for NGS. Pool and clean up.
Day 2
Sequence.
Analyze counts, fit ladder, produce target by bin matrix, call proteoforms.
Review acceptance criteria.
19. Calculations you will use
dPCR occupancy. λ = −ln(1 − k/N). Molecules per bin = λ·N.
Per‑target correction. Molecules corrected = molecules measured divided by ρ.
Ladder mapping. Fit log10(MW) = a + b·t + c·t^2 on ladder peaks. Then MW(bin j) = 10^(a + b·tj + c·tj^2).
20. Buffer recipes
PBS, pH 7.4. 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4.
HEPES buffer, pH 7.5. 50 mM HEPES, 150 mM NaCl.
Bead capture buffer. PBS, 0.05 percent Tween‑20, 1 mg mL‑1 BSA.
Wash buffer. PBS, 0.05 percent Tween‑20.
Collection buffer. 20 mM HEPES pH 7.5, 150 mM NaCl, 0.05 percent Tween‑20, 10 mM methyl‑beta‑cyclodextrin, 2 mg mL‑1 BSA.
Cleavage buffer example for disulfide. PBS with 25 mM TCEP, 10 minutes at room temperature.
21. Minimum documentation set
Run log with injection time, separation field, bin timing.
Ladder fit coefficients and residuals.
Detector lot QC with 1:1 stoichiometry evidence.
Per‑target ρ values and dates.
Raw bin counts, UMI collapse stats, normalization factors, and final matrices.
Acceptance criteria outcomes and any deviations.
Final notes
Use NGS readout for full DSI‑Seq because it solves bin indexing cleanly and scales to 24 or more targets. Keep dPCR for targeted troubleshooting or a very small panel.
The most common failure in early runs is residual SDS killing binding. Your collection buffer and bead wash are the leverage points.
Keep the chemistry protein‑only by continuing to nuclease treat lysates and by ensuring the only DNA that can be amplified comes from the single‑tag detector after cleavage.
This is written to let you prototype, optimize, and validate the method on benchtop equipment. The protocol implements the IP‑lean single‑tag digital immunoPCR style readout inside each size bin, and it avoids proximity ligation or proximity extension chemistry. It supports two readouts:
NGS counting for high multiplex and practical bin indexing.
dPCR counting for small target panels or early debugging.
The workflow is plate‑based, since that is the fastest and most accessible path to working data.
0. Scope and outcome
Goal. Resolve proteins by apparent size into many narrow fractions, then quantify specific targets in each fraction by converting each captured protein into one DNA tag and counting tags. Output is a matrix of counts: targets by size bins, calibrated to kDa using a co‑run ladder.
Typical first build. 48 bins across a 12 minute separation window, 12 to 24 targets by NGS readout, input 1 to 10 micrograms total protein per lane, results in one day after library prep.
1. Safety and handling
Acrylamide and SDS are irritants. Handle with gloves, lab coat, and eye protection. Dispose of acrylamide waste as hazardous.
UV photocleavage variants require eye and skin protection. Shield the UV source.
Nucleases can aerosolize. Open tubes slowly and decontaminate surfaces.
Segregate pre‑PCR and post‑PCR spaces. Use UDG carryover control.
2. Materials and reagents
2.1 Buffers and solutions
Lysis buffer, PCR compatible50 mM HEPES pH 7.5, 150 mM NaCl, 1 percent NP‑40 or 0.5 percent Triton X‑100, 10 percent glycerol, protease inhibitors, phosphatase inhibitors if needed.
Optional SDS denaturing load buffer for chip1x TGS or TBE, 0.1 to 0.5 percent SDS, 10 mM DTT, 10 percent glycerol. Heat 70 to 90 C for 5 minutes if full denaturation is required.
Replaceable sieving polymer for SDS‑CECommercial SDS‑CE polymer or 2.5 to 4.0 percent linear polyacrylamide in 1x TBE with 0.1 percent SDS.
Collection buffer per well20 mM HEPES pH 7.5, 150 mM NaCl, 0.05 percent Tween‑20, 10 mM methyl‑beta‑cyclodextrin, 2 mg mL‑1 BSA. The cyclodextrin scavenges SDS on contact.
Bead capture bufferPBS, 0.05 percent Tween‑20, 1 mg mL‑1 BSA.
Wash bufferPBS, 0.05 percent Tween‑20.
Cleavage bufferDepends on linker. For disulfide linkers: PBS with 10 to 50 mM TCEP. For o‑nitrobenzyl photocleavage: PBS.
UDG mix for carryover controldNTPs with dUTP substitution, thermolabile UDG.
Benzonase mix25 to 50 U mL‑1 Benzonase with 2 mM MgCl2. Stop with 10 mM EDTA or heat inactivation per supplier.
2.2 Controls and standards
Protein ladder 10 to 250 kDa, prestained.
Spike‑in protein standards for at least two targets, quantified concentration.
Negative control lysate with phosphatase treatment for PTM assays.
2.3 Antibodies and DNA tags
Capture antibodies coupled to magnetic beadsUse Protein G beads with oriented coupling or covalent coupling kits. Aim for 1 to 5 micrograms antibody per mg beads. Prepare one capture bead type per target.
Detector binders that are monovalent with one DNA tag eachPrefer Fab fragments or nanobodies engineered with a single cysteine. Conjugate one DNA tag per binder by maleimide‑thiol chemistry. Confirm 1:1 stoichiometry by intact mass.
DNA tag designLength 60 to 100 nt.Layout: 5’ universal priming site A, 8 to 12 nt UMI, 12 to 16 nt target ID, 5’ half of an Illumina adapter if using NGS, 3’ blocking group if needed.The tag is attached to the detector via a generic, cleavable linker. No two‑probe hybridization is used.
Per‑bin index adapters for NGSShort double‑stranded adapters with bin‑specific index sequences and universal priming sites. These are ligated or appended in PCR to encode bin identity.
TaqMan probes for dPCROne probe per target tag. For dPCR you cannot multiplex many bins in one tube, so plan per‑bin reactions only for 1 to 4 targets.
2.4 Equipment
Microchip or capillary SDS electrophoresis instrument with replaceable polymer.
Simple fraction collector setupA motorized XY stage for a 96‑well plate under the capillary outlet. Alternatively a commercial fraction collector with programmable step time.
Magnetic racks for 96‑well plates.
Thermocycler and a bench NGS library prep kit.
dPCR instrument if using the dPCR readout.
Plate shaker, microcentrifuge, fluorometer or qPCR for library QC.
Optional UV source for photocleavage, with shielding.
3. Panel design and conjugation
Choose targets. Begin with 12 to 24 proteins. Favor clones validated for Western or IP. Include one reference housekeeper.
Epitope check. Select capture and detector epitopes that do not compete. For oligomers or large complexes, expect a calibration factor as noted later.
Conjugate detectors.
Reduce Fab or nanobody to expose the engineered cysteine.
React with maleimide‑tagged DNA at 1.5 to 2.0 molar excess.
Purify by size exclusion or affinity to remove free DNA.
Confirm 1:1 by intact mass or denaturing CE.
Bead coupling. Couple capture antibody to magnetic beads. Block with BSA. Store in PBS with 0.02 percent sodium azide at 4 C.
Document the per‑target conversion factor ρ. If two detectors can bind one captured protein or the protein is oligomeric, measure ρ using purified protein and record it in the target sheet.
4. Sample preparation
Harvest cells or tissue. Keep cold.
Lyse in PCR‑compatible lysis buffer. For maximum size resolution across proteoforms you may choose an SDS denaturing load immediately before separation.
Clarify by spinning at 14,000 g for 10 minutes.
Protein quant by BCA.
Nuclease treatment for protein‑only readout. Add Benzonase mix to lysate, incubate 15 minutes at room temperature. Stop with EDTA or heat inactivate. This ensures no endogenous DNA or RNA is counted.
Spike‑in controls. Add defined copies of two recombinant controls.
Add ladder for a dedicated ladder run. Prepare a separate tube with the prestained ladder in the same matrix for calibration runs.
5. Microchip SDS separation
You can run native or denaturing. For size indexing you will typically run SDS.
Prime the chip with replaceable sieving polymer per instrument instructions.
Equilibrate with 1x running buffer containing 0.1 percent SDS.
Load sample. If using SDS, mix lysate with SDS load buffer to 0.1 to 0.5 percent SDS and heat 70 to 90 C for 5 minutes.
Injection. Electrokinetic inject for 3 to 5 seconds at reduced field.
Separation. Field strength 200 to 300 V cm‑1, total separation window 10 to 15 minutes. Record the exact time zero when injection ends.
Run a separate ladder trace with the same separation program and collect fractions the same way as the sample run.
6. Plate‑based fractionation into size bins
Prepare a 96‑well plate. Dispense 10 to 15 microliters of collection buffer into the wells you will use.
Define bins. For a 12 minute separation choose 48 bins at 15 seconds per bin. That uses half a 96‑well plate.
Position the capillary outlet above the first well. Use a droplet‑friendly tip or align to touch the meniscus.
Collect. Start separation and move the plate on a schedule so that each bin collects exactly 15 seconds of eluent. A simple script on the stage controller is enough.
Finish. Seal the plate. The cyclodextrin in the collection buffer reduces free SDS below 0.02 percent in seconds and helps renaturation of linear epitopes.
Optional fraction concentration. If fractions are very dilute, you can add 1 mg of dry Bio‑Beads SM‑2 per well for 5 minutes to further scavenge SDS, then remove beads with a magnetic wand or by decanting. Validate that your targets are not lost.
7. Per‑bin immunoassay with single‑tag detectors
This yields one DNA tag per captured protein complex in each bin. No proximity ligation or extension is used.
For each bin well:
Add capture beads
Add 10 to 20 microliters of bead suspension with the capture antibody for target i, or use a bead mix containing distinct bead codes if you run targets in parallel within a bin.
Incubate 30 minutes at room temperature with gentle shaking.
Magnet and wash
Pull beads down. Remove supernatant.
Wash 3 times with 100 microliters wash buffer.
Detector binding
Add the monovalent detector conjugate carrying a single DNA tag, at 1 to 5 nM in bead capture buffer.
Incubate 30 minutes.
Wash 3 times.
Release DNA tags
Add 20 microliters cleavage buffer.
For disulfide linkers, incubate with 10 to 50 mM TCEP for 10 minutes.
For photocleavage, illuminate with the specified UV for the vendor’s o‑nitrobenzyl linker. Keep temperature controlled.
Collect the supernatant which contains the released DNA tags.
UDG carryover control
If you use dUTP in tags, add thermolabile UDG and incubate per supplier before amplification to remove any carryover products. The released tags do not contain prior amplicons.
Pooling strategy
For NGS readout keep each bin separate through the next indexing step.
For dPCR readout proceed per bin per target. dPCR is best limited to 1 to 4 targets across a small number of bins during development.
Notes
If you prefer, perform steps 1 to 3 in bulk per bin with a mixed bead cocktail for all targets. This speeds handling. The detector step remains one DNA per bound protein because each detector is monovalent.
If an antigen accepts two detectors per captured protein, apply the per‑target factor ρ during analysis.
8. Readout A. NGS library and sequencing
NGS is the practical route for many bins and many targets because you can encode bin identity by indexing.
Per‑bin index addition
Set up a PCR using primers that add a unique bin index i5 or i7 to each bin. 10 to 14 cycles are typical since the tag pool can be low copy.
Use a common reverse primer so that only the index changes across bins.
Target identity in the tag
The target ID is already encoded in the tag sequence. You do not add any probe hybridization step between oligos on two antibodies. You simply amplify what you cleaved.
Pool and clean up
Pool equal volumes or equal mass from all bin PCRs for a sample.
Clean up with SPRI beads at 1.2x ratio.
QC and run
Measure library size and concentration. Expect 150 to 300 bp.
Sequence paired‑end or single‑end, 50 to 100 cycles is usually enough.
Depth target per sample 1 to 3 million reads for a 24 target by 48 bin design.
Demultiplex
Demultiplex by sample and by bin index.
Parse reads to tag IDs and UMIs. Collapse UMIs to unique molecules.
9. Readout B. dPCR per bin for 1 to 4 targets
Use this for early method checkout or when you only need a few targets.
Set up dPCR reactions
For each bin and each target, set up a reaction with the target‑specific TaqMan probe.
Partition and run endpoint PCR.
Count
Record positive partitions k and total partitions N.
Compute lambda per bin as λ = −ln(1 − k/N).
Molecules per bin equals λ times N.
Scale
Apply the per‑target calibration factor ρ if relevant.
Continue to the size calibration step below.
10. Size calibration using the ladder
You will convert bin index to apparent kDa.
Collect a ladder run with identical fractionation
Run the prestained ladder alone with the same separation and fraction schedule used for the sample. Collect into a plate with the same 48 bins.
Quantify ladder protein abundance in each bin by simple absorbance at 595 nm if dye permits, or by a quick fluorometric protein assay across wells.
Identify peaks
Ladder proteins will appear as peaks across bins. Fit peak centers to their known molecular weights.
Fit the mapping
A robust model is: log10(MW) = a + b·t + c·t^2 where t is bin center time in minutes.
Fit a, b, c by least squares using the ladder peaks.
For each bin j in the sample, compute its center time and then its apparent kDa by inverting the relationship.
Uncertainty
Compute 95 percent confidence intervals on MW per bin from the residuals of the fit.
11. Data processing and outputs
Counts table
For NGS, produce a table of unique molecule counts per target per bin after UMI collapse.
For dPCR, produce molecules per bin from Poisson correction.
Normalize
Divide by the external spike‑in recovery to correct for run‑to‑run variation.
Optionally normalize per bin by a total protein signal if measured.
Proteoform calling
For each target, smooth the binned profile with a small window.
Identify local maxima as proteoform peaks.
Report peak bin, apparent kDa, and integrated counts under each peak.
Final report
Matrix: targets × bins with counts.
Proteoform summary table with apparent kDa, counts, fraction of total per target.
QC panel with ladder fit, spike‑in recovery, background levels, replicate CVs.
12. Acceptance criteria and QC
Adopt these hard thresholds before you trust a run.
Reagent‑only background. After full library prep with no protein, fewer than 0.2 percent of reads assign to valid tag IDs or fewer than 0.2 positives per 20,000 partitions for dPCR.
Blank lysate after nuclease. With universal primers but no released tags, signal must be at background.
Spike‑in recovery. 0.8 to 1.2 across a 100‑fold range.
Dilution linearity. Two dilutions at 2x load must yield 1.8 to 2.2x counts per bin.
Ladder fit residual. Standard error of log10(MW) fit less than 0.03.
Replicates. Technical replicate CV less than 10 percent in bins with at least 400 molecules.
13. Validation plan
Run in this sequence.
Separation and fractionation stub
Ladder only. Build the bin to kDa mapping and verify timing precision over three runs.
Single‑target end‑to‑end
One target in a control lysate. Spike a dilution series of the purified protein. Validate linearity and the expected single peak at the known kDa.
Isoform resolution
Use a biology that creates a known cleavage, for example PARP1 cleavage in apoptosis. Confirm two peaks at expected apparent sizes.
PTM specificity
Use a phospho target with a modification specific detector. Compare stimulated vs phosphatase treated. The modified target should vanish in treated samples without a shift in the pan profile.
Multiplex cross‑talk
Build the panel in subpanels of 6 to 8 targets. Check background in no‑antigen wells and confirm no off‑target rises when subpanels are combined.
Reproducibility and lot stability
Ten technical replicates. Track per‑bin per‑target CV.
New detector lots get a check of the conversion factor ρ and a small ladder redo.
14. Troubleshooting
Weak or no signal in all binsCheck detector conjugation efficiency and cleavage step. Confirm 1:1 stoichiometry. Verify that the UDG step is not destroying tags.
High background in no‑protein controlsSuspect amplicon carryover or free DNA tag contamination. Increase physical separation of spaces, add UDG, and re‑purify detectors to remove free tag.
Flat size profile without peaksFraction timing off or excessive diffusion. Shorten bin width to 10 seconds or increase field strength. Confirm collection buffer is dispensed before the run.
Shifted ladder mapping between runsTemperature or polymer batch differences. Always run a ladder with each batch and refit a, b, c. Apply mapping per run.
Loss of antibody binding due to SDSIncrease cyclodextrin concentration in collection buffer to 20 mM. Add a short bead‑based buffer exchange before detector binding. Use clones validated for linear epitopes.
15. Options and upgrades
Droplet fractionationReplace the plate with a T‑junction droplet maker that encapsulates the outlet stream at a fixed rate. A side stream can inject a droplet index oligo that is appended later by standard ligation. This increases bin count to 96 to 128 at the same run time.
Bin reduction mergeFor abundant targets, merge adjacent bins in software to increase counts per bin and improve precision.
Alternative SDS scavengersPotassium chloride can precipitate SDS as potassium dodecyl sulfate. Use with care due to protein precipitation. Validate recovery on spike‑ins.
16. IP and compliance posture
One DNA tag per detector binder.
No proximity ligation or proximity extension between two nucleic acid‑bearing probes.
No single‑molecule enzyme arrays.
Bin identity is appended during library prep by standard indexing, or dPCR is run per bin per target without any probe‑probe interaction.
Include UDG carryover suppression and commodity linkers only.
Record these choices in your design history file from day one.
17. Starter T cell panel
Begin with 12 to 18 targets that are Western or IP validated and bind linear epitopes. For example: CD3ζ, ZAP70, LAT, SLP76, PLCG1, ERK1, ERK2, AKT, mTOR, 4EBP1, NF‑κB p65, PARP1, and a reference such as beta‑actin. Add phospho‑specific detectors as a separate subpanel and always run a phosphatase control.
18. Example day plan
Day 1 morning
Conjugate 2 to 4 detectors and QC one by mass.
Couple 2 to 4 capture antibodies to beads and block.
Day 1 afternoon
Prepare lysates, nuclease treat, spike controls.
Run ladder and sample separations with fractionation to 48 bins.
Day 1 evening
Perform per‑bin bead capture, detector binding, cleavage.
Start per‑bin index PCR for NGS. Pool and clean up.
Day 2
Sequence.
Analyze counts, fit ladder, produce target by bin matrix, call proteoforms.
Review acceptance criteria.
19. Calculations you will use
dPCR occupancy. λ = −ln(1 − k/N). Molecules per bin = λ·N.
Per‑target correction. Molecules corrected = molecules measured divided by ρ.
Ladder mapping. Fit log10(MW) = a + b·t + c·t^2 on ladder peaks. Then MW(bin j) = 10^(a + b·tj + c·tj^2).
20. Buffer recipes
PBS, pH 7.4. 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4.
HEPES buffer, pH 7.5. 50 mM HEPES, 150 mM NaCl.
Bead capture buffer. PBS, 0.05 percent Tween‑20, 1 mg mL‑1 BSA.
Wash buffer. PBS, 0.05 percent Tween‑20.
Collection buffer. 20 mM HEPES pH 7.5, 150 mM NaCl, 0.05 percent Tween‑20, 10 mM methyl‑beta‑cyclodextrin, 2 mg mL‑1 BSA.
Cleavage buffer example for disulfide. PBS with 25 mM TCEP, 10 minutes at room temperature.
21. Minimum documentation set
Run log with injection time, separation field, bin timing.
Ladder fit coefficients and residuals.
Detector lot QC with 1:1 stoichiometry evidence.
Per‑target ρ values and dates.
Raw bin counts, UMI collapse stats, normalization factors, and final matrices.
Acceptance criteria outcomes and any deviations.
Final notes
Use NGS readout for full DSI‑Seq because it solves bin indexing cleanly and scales to 24 or more targets. Keep dPCR for targeted troubleshooting or a very small panel.
The most common failure in early runs is residual SDS killing binding. Your collection buffer and bead wash are the leverage points.
Keep the chemistry protein‑only by continuing to nuclease treat lysates and by ensuring the only DNA that can be amplified comes from the single‑tag detector after cleavage.
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