About

Vladislav (Vlad) Belyy is an Assistant Professor at The Ohio State University in the Department of Chemistry and Biochemistry, beginning his independent research career in August 2022. His research focuses on understanding how cells relay information across membranes, particularly through the study of oligomerization of signaling proteins. His lab develops precise and quantitative approaches to perturb and measure oligomerization in native environments, utilizing techniques such as single-molecule imaging, protein engineering, genome editing, in vitro biochemistry, and quantitative microscopy. Belyy's work aims to reveal new insights into stress sensing by cells and to provide tools for studying signaling proteins in cell biology.

Research topics

• Biophysics
• Chemistry
• Cell biology
• Biology
• Biochemistry

Selected publications

• BPS2026 – Investigating activity of Eph receptors with direct control over oligomerization

  Biophysical Journal · 2026-02-01

  articleSenior author
  PublisherDOI

• BPS2026 – Dissecting the activation mechanism of the bi-functional kinase/RNase IRE1 with orthogonal control over oligomerization and phosphorylation

  Biophysical Journal · 2026-02-01

  articleSenior author
  PublisherDOI

• BPS2026 – Probing transcriptional and splicing regulation in the unfolded protein response with optically activated IRE1

  Biophysical Journal · 2026-02-01

  articleSenior author
  PublisherDOI

• BPS2026 – Intricate regulation of RNA cleavage and splicing by the endoribonuclease IRE1

  Biophysical Journal · 2026-02-01

  articleSenior author
  PublisherDOI

• Data and analysis code for the manuscript titled "Direct Optical Activation of Human IRE1 Identifies Unique Patterns of Transcriptional and Post-Transcriptional mRNA Regulation in the Unfolded Protein Response""

  Zenodo (CERN European Organization for Nuclear Research) · 2025-12-18

  otherOpen access1st authorCorresponding

  This repository contains the raw data and analysis code for our manuscript (Smith et. al.), tittled "Direct Optical Activation of Human IRE1 Identifies Unique Patterns of Transcriptional and Post-Transcriptional mRNA Regulation in the Unfolded Protein Response".

  PublisherDOI

• Optogenetic Clustering of Human IRE1 Reveals Differential Regulation of Transcription and mRNA Splice Isoform Abundance by the UPR

  bioRxiv (Cold Spring Harbor Laboratory) · 2025-07-21

  preprintOpen accessSenior authorCorresponding

  Inositol-requiring enzyme 1 (IRE1) is one of three known sensor proteins that respond to homeostatic perturbations in the metazoan endoplasmic reticulum. The three sensors collectively initiate an intertwined signaling network called the Unfolded Protein Response (UPR). Although IRE1 plays pivotal roles in human health and development, understanding its specific contributions to the UPR remains a challenge due to signaling crosstalk from the other two stress sensors. To overcome this problem, we engineered a light-activatable version of IRE1 and probed the transcriptomic effects of IRE1 activity in isolation from the other branches of the UPR. We demonstrate that 1) oligomerization alone is sufficient to activate IRE1 in human cells, 2) IRE1's transcriptional response evolves substantially under prolonged activation, and 3) the UPR induces major changes in mRNA splice isoform abundance in an IRE1-independent manner. Our data reveal previously unknown targets of IRE1 transcriptional regulation and direct degradation. Additionally, the tools developed here will be broadly applicable for precise dissection of signaling networks in diverse cell types, tissues, and organisms.

  PublisherOA PDFDOI

• Direct optical activation of human IRE1 identifies unique patterns of transcriptional and post-transcriptional mRNA regulation in the unfolded protein response

  Journal of Biological Chemistry · 2025-12-18

  articleOpen accessSenior author

  Inositol-requiring enzyme 1 (IRE1) is one of the three known sensor proteins that respond to homeostatic perturbations in the metazoan endoplasmic reticulum. The three sensors collectively initiate an intertwined signaling network called the unfolded protein response (UPR). Although IRE1 plays pivotal roles in human health and development, understanding its specific contributions to the UPR remains a challenge due to signaling crosstalk from the other two stress sensors. To overcome this problem, we engineered a light-activatable version of IRE1 and probed the transcriptomic effects of IRE1 activity in isolation from the other branches of the UPR. We demonstrate that 1) oligomerization alone is sufficient to activate IRE1 in human cells, 2) IRE1's transcriptional response evolves substantially under prolonged activation, and 3) the UPR induces major changes in mRNA splice isoform abundance in an IRE1-independent manner. Our data reveal previously unknown targets of IRE1's transcriptional regulation and direct degradation. Additionally, the tools developed here will be broadly applicable for precise dissection of the UPR in diverse cell types, tissues, and organisms.

  PublisherOA PDFDOI

• Optogenetic platform for dissecting IRE1's enzymatic mechanism

  Biophysical Journal · 2024-02-01

  articleSenior author
  PublisherDOI

• Dissecting ER stress signaling with live-cell single-molecule imaging and optogenetics

  Biophysical Journal · 2024-02-01

  articleOpen accessSenior author
  PublisherOA PDFDOI

• Endogenous tagging of IRE1 in U-2 OS cells

  BIO-PROTOCOL · 2023-01-01

  articleOpen accessSenior author

    Overview This protocol describes the insertion of a C-terminal tag (specifically, HaloTag) into the endogenous locus of the human ERN1 gene, which encodes the ER stress sensing protein IRE1α. It is important to note that, while this protocol reflects the procedure exactly as we performed it1, it is not a well-optimized protocol and results in very low editing efficiency. For someone seeking to introduce an endogenous tag into the ERN1 locus in a different cell line, we strongly recommend combining the useful features of this protocol (e.g. guide RNA sequence and homology directed repair (HDR) template sequence) with a more efficient strategy for CRISPR/Cas9-directed genome editing. For instance, our colleagues recently achieved substantially higher editing efficiency by pre-treating cells with nocodazole and nucleofecting a pre- assembled Cas9-sgRNA complex into the cells together with the HDR template DNA2. In contrast, our approach relied on a transient transfection of the target cells with two plasmids, one encoding both the Cas9 protein and the sgRNA and the other carrying the HDR template sequence, largely as described in part 3 of the book chapter by K. L. McKinley3. While this approach is simpler, it produced such low editing efficiencies that we were unable to modify every ERN1 allele in U-2 OS cells, which are hypertriploid (according both to our own observations and to the American Type Culture Collection, ATCC). Instead, we started with a clonal population of U-2 OS cells in which all but one ERN1 alleles had been knocked out via CRISPR/Cas9-directed editing of exon 1. This partial IRE1α knockout cell line was generated as a by-product of our earlier effort to completely knock out IRE1α in U-2 OS cells4. The logic behind using a partial knock-out is that only one allele needs to be edited in order to ensure that all IRE1α protein contains the HaloTag. In summary, while this protocol worked for us, it was inefficient and time-consuming, and we would like to be transparent about the fact that there exist better alternatives to achieve endogenous tagging of the ERN1 locus in human cells.  Protocol Part 1: Selection and preparation of parental cells. As discussed in the Overview section, U-2 OS cells (and many other commonly used immortalized cell lines) are hypertriploid. The low efficiency of this protocol means that the chances of finding a clone in which every allele carries the desired C-terminal insertion is very low. One way to resolve this issue is to start with a clonal population of partial knock-out cells that carry only a single intact copy of the ERN1 gene. Such partial knock-out cells will typically get generated as a byproduct of any attempt to generate a CRISPR knock-out cell line, since CRISPR editing is not perfectly efficient and every CRISPR round will result in a mixed population of fully edited, partially edited, and unedited cells. For the details on generating IRE1α knock-out U-2 OS cells in house, please refer to our earlier paper4. Alternatively, a mixed population of partial and full knock-out cells can be obtained commercially from a company such as Synthego. The following steps describe the identification of a clonal population of a cell containing only a single intact ERN1 locus. The approach has been directly applied to U-2 OS cells but should work with minimal modifications for most immortalized adherent cell lines. 1. Separate the mixed polyclonal population of edited cells into single clones. There are two ways of getting individual cell clones: limiting dilution and plate sorting. Note: unless specified otherwise, cells are grown in fully supplemented growth medium: high glucose DMEM (e.g. Thermo Fisher Cat# 11965092), 10% fetal bovine serum (FBS), 2 mM L-glutamine, 50 I.U./mL of penicillin-streptomycin. 1.1 Option 1: Limiting Dilution 1.1.1  Grow the mixed cell population in a well of a 6-well plate or on a 10 cm dish until the cells are ~70% confluent.1.1.2  Dissociate and count the cells (we recommend counting in triplicate and taking the average of the measurements, since an accurate count is very important for the subsequent step).1.1.3  Calculate the dilution needed to plate the cells into a 96 well plate with a final density of 0.5 cells/well. At the recommended volume of 150 µl of growth media per well, this corresponds to ~3.3 cells/mL.1.1.4  Dilute the cells as calculated above and seed them into wells of four 96-well plates (384 wells total). Note #1: this step can be adjusted to use more or fewer plates if desired, trading off the required resources for the probability of finding good clones. Note #2: as a backup for unexpectedly low cell viability after limiting dilutions, we recommend also plating 1 or 2 96-well plates at a higher calculated seeding density of 2 cells per well. 1.2 Option 2: Single-cell Plate Sort. This might be a better way of getting clones, but our fluorescence-activated cell sorting (FACS) machine was malfunctioning at the time of the experiment so we were not able to get plate sorting to work reliably. We used option 1, limiting dilution, instead. 1.2.1  Grow cells on a 15 cm dish or in a T150 flask until ~70% confluent.1.2.1  Trypsinize the cells, quench the trypsin with an equal volume of fresh growth medium, and pellet the cells by gentle centrifugation (300 RCF for 3 minutes).1.2.3  Resuspend the pellet in FACS medium (for U-2 OS, PBS supplemented with 0.5% FBS seems to work fine).1.2.4  Place the cells on ice.1.2.5  As quickly as possible, perform a plate sort, targeting one cell into every well of a 96-well plate. Aim to seed 3-5 plates since you won't know at the start what your survival rate will be. The exact protocol for the plate sort will depend on your specific FACS machine. In terms of gating, the cells are not expressing any fluorescent markers yet, so just use forward and side scatter gates to remove debris and cell clumps, allowing any singlet cells through. 1.3 Recover individual clones. (Regardless of whether you picked Option 1 or Option 2 above). 1.3.1  Return the 96-well plates to the incubator and replace the growth medium every 3-4 days. Approximately 1.5 weeks after seeding, start scanning through the wells using a standard phase contrast microscope to identify wells that have healthy growing clones. A good clone will appear as a single, roughly circular tight clump of cells. Take care to only select wells with a single clone: if a well has two or more separate cell clumps, it should not be used in subsequent steps since it will contain a polyclonal mixture of cells. Note: you may not be able to find all the good clones at the 1.5 week mark since cells will take variable amounts of time to recover from a single-cell stage. Keep checking the plates for at least another 1-2 weeks and taking notes of any newly identified clones.1.3.2  As some of the clones start getting dense in the wells of a 96-well plate (i.e. the clump of cells begins to occupy a significant fraction of the well’s surface area), split each clone into two wells of a 24-well plate. One well will be used for expanding the cells, and the other for genomic DNA (gDNA) analysis. Note: due to the variable growth rates, this process will not be perfectly synchronized for all clones. Process the individual clones as they get ready. 2. Screen the clones to identify ones with a single intact ERN1 allele. 2.1  Once any cells grow to near-confluency in a well of the 24-well plate reserved for gDNA analysis, harvest it and purify the gDNA using your protocol of choice. We recommend the PureLink Genomic DNA Mini Kit (Thermo Fisher, cat# K182002). If using the recommended kit, follow the instructions exactly and elute the purified gDNA in 25 µl of 2 mM Tris buffer, pH 8.0.2.2  Measure the concentration of the purified gDNA on a Nanodrop instrument or using your method of choice; your yield should be at least 10 ng/µl for a successful subsequent PCR reaction (preferably above 30 ng/µl).2.3  Prepare PCR reactions for each gDNA sample to amplify the edited regions as follows: ReagentAmount5XPhusion GC Buffer (New England BioLabs Cat# B0519S)                                                                   5 µLdNTP mix (10 mM each)0.5 µLDMSO0.75 µLForward Primer, 10 µM (5'-AGCGCTTATAGGGCCGGGAA-3')1.25 µLReverse Primer, 10 µM (5'- GTTCAAACAAGGATTCGAAGCGCAGG-3')1.25 µLPhusion DNA Polymerase (New England BioLabs Cat# M0530S)0.5 µLTemplate DNA (concentration measured by NanoDrop)Sufficient volume to give 100 ng ofgDNA Nuclease-free waterSufficient volume to bring total reaction volume up to 25 µL 2.4  Amplify the reactions in a standard PCR cycler using the following program: Step #DescriptionTemperatureDuration (min:sec)1Initial denaturation95°C5:002Denaturation98°C0:203Annealing69°C0:154Extension72°C0:30 Go to step 2 (repeat 31x)  5Final extension72°C3:006Sample ready10°Chold 2.5  Clean up the PCR products using a method of your choice (we use the DNA Clean & Concentrator-5 kit, Zymo Research Cat# D4014, following the manufacturer’s instructions) and measure the resulting DNA concentration. You should elute the gDNA in 10 µL of elution buffer and get a yield of at least 500 ng total DNA (> 50 ng/µL). NOTE: it may be a good idea to run the PCR product on a gel to ensure that you are getting a single strong band at the expected size. If you are not, you’ll want to troubleshoot your PCR first before proceeding with TOPO cloning.2.6  Clone the PCR product into a TOPO vector (we use the Zero Blunt TOPO PCR Cloning Kit, Thermo Fisher Cat# 451245). Note: The reason for doing TOPO cloning is to get individual sequencing results for every allele in a given clone. Remember that the master PCR product you generated above will contain a mixture of every allele; by cloning it into TOPO vectors, transforming the vectors into bacterial cells, and sequencing many colonies for each clone, you can get individual insertion/deletion sequences for each of the alleles. 2.6.1  Assemble the TOPO cloning reactions (one for each PCR product) as follows: ReagentAmountPCR product (~50-100 ng/ µL)1 µLSalt solution (included with kit)0.5 µLWater1 µLpCR™Blunt II-TOPO™ mix0.5 µL 2.6.2  Mix gently and incubate for 5 minutes at room temperature.2.6.3  Place the reaction on ice and proceed immediately to the next step (transformation). 2.7  Transform the TOPO reactions into competent E. coli cells of your choice. We transformed 2 µL of the TOPO mixture into 16 µL of Stellar chemically competent cells (Takara Bio Cat# 636766) following the manufacturer’s instructions, but you may need to modify the amounts if using different competent cells).2.8  Plate the cells on pre-warmed LB-Kanamycin plates and wait for colonies to grow overnight.2.9  Perform colony PCRs on ~30 individual colonies for each gDNA sample: 2.9.1  Prepare a PCR master mix for all reactions as follows: ReagentAmount per reaction5XPhusion HF Buffer (New England BioLabs Cat# B0518S)3 µLdNTP mix (10 mM each)0.4 µLForward Primer, 18 µM (M13 Forward [−20], 5'- GTAAAACGACGGCCAG -3')0.25 µLReverse Primer, 18 µM (M13 Reverse,5'- CAGGAAACAGCTATGAC-3')0.25 µLPhusion DNA Polymerase (New England BioLabs Cat# M0530S)0.1 µLNuclease-free water10.9 µL 2.9.2  Split the master mix into as many PCR tubes as needed, pipetting 15 µL into each tube.2.9.3  Add bacterial cells from a different colony to each PCR tube. To do this, just barely touch the colony with a pipette tip and dip it into the PCR tube, pipetting up and down a couple times to mix. You only need a tiny number of cells; too many cells can cause the PCR to fail.2.9.4  Amplify the reactions in a standard PCR cycler using the following program:  Step #DescriptionTemperatureDuration (min:sec)1Initial denaturation98°C5:302Denaturation98°C0:103Annealing52°C0:304Extension72°C0:30 Go to step 2 (repeat 32x)  5Final extension72°C10:006Sample ready10°Chold 2.9.5  (optional but recommended) Run the colony PCR products on a large 1% agarose gel. Colony PCR can be finicky, and it's a good idea to check which reactions have a good product (you expect to see a sharp single band in the 700-800 bp size range) before wasting money on sequencing reactions. 2.10  Send the PCR reactions that showed a clear band out for Sanger sequencing with a compatible primer (e.g. M13 Forward [-21], 5′-GTA AAA CGA CGG CCA GT-3′).2.11  Once the sequencing results are in, align them to the sequence of human ERN1 exon 1. If you have a sufficient number of clones, you should be able to see a clear pattern, where every modified allele shows up multiple times (see examples of multiple identical edited sequences from a single clone of mammalian cells).  2.12  You may need to send out a few more bacterial colony PCRs for sequencing as needed, but once you have enough sequencing data, you should be able to identify a good clone that contains a single intact (WT) allele. To estimate allele copy number, compare the number of sequencing reads for each modification 2.12.1  For example: if you have allele versions A, B, C, and D, and you get 8 reads for allele A, 3 reads for B, 4 for C, and 3 for D, your clone likely contains two alleles of allele A and one copy each of B, C, and D. This is not an exact approach but given enough sequencing data it should be reasonably accurate. 2.13  To move forward with the protocol, you'll need to identify a clone that contains frameshifts or stop codons in every ERN1 allele except for one unedited (WT) allele. To be extra sure, it may be a good idea to check IRE1 protein expression levels and stress-induced phosphorylation in the selected clone(s) by Western blotting before proceeding.  Part 2: Insertion and validation of the tag into the chosen parental cells. 3. Insert a C-terminal HaloTag into the chosen clonal population by CRISPR/Cas9 genome editing. 3.1  Count and seed your chosen cells from the previous section for transfection (80,000 cells into one well of a 6-well plate).3.2  Grow overnight cultures of the two plasmids required for transfection: pPW3754, which contains the SpCas9 protein and the sgRNA targeting the C-terminus of human IRE1α, and pPW3755, which contains the HDR template for inserting a C-terminal HaloTag.3.3  Purify the plasmid DNA from the two overnight cultures, making sure to use a minprep kit that generates endotoxin-free DNA suitable for transient transfections. Note: we found that using freshly miniprepped plasmid DNA (a few weeks old or less) helps boost transfection efficiency.3.4  Two days after the human cells were seeded into the well of a 6-well plate, they should be ~60% confluent and ready for transfection. Transfect them as follows: 3.4.1  Mix 1.6 µg of pPW3754 plasmid DNA and 1.6 µg of pPW3755 plasmid DNA in a 15 mL conical tube.3.4.2  Add enough room-temperature Opti-MEM I reduced serum medium (Thermo Fisher Cat# 31985062) to the tube to bring the total volume of the DNA + OptiMem mixture to 140 µL (e.g. If you added 3 µL of pPW3754 and 6 µL of pPW3755, add 131 µL of OptiMem). Mix well.3.4.3  Add 11 µL of Fugene HD transfection reagent (Promega Cat# E2311) directly to the DNA- OptiMem solution and immediately vortex the tube for 5 seconds to mix.3.4.4  Incubate the solution at room temperature for 5 minutes (not longer!).3.4.5  Aspirate the growth medium from cells.3.4.6  Quickly 3 mL of pre-warmed fully supplemented medium (containing 10% FBS and 2 mM L-glutamine) but without Penicillin/Streptomycin to the conical tube containing the DNA- Fugene mixture and mix by pipetting up and down a couple of times.3.4.7 Gently add the media with DNA and Fugene to the cells. There is no need to remove the transfection reagent after several hours. 3.5  1-2 days post transfection (once the cells get ~90% confluent), split them into a 10 cm dish.3.6  Once the 10 cm dish is ~70% confluent, split it into one 10 cm dish for maintenance and one 15 cm dish (or T150 flask) for FACS. As soon as you have enough cells in the maintenance dish, we recommend freezing a couple of aliquots as “checkpoints” before expanding the cells further. 4. Isolate and identify correctly edited clones. 4.1  Once the cells in the FACS flask are ~70% confluent, they should be ready to sort. In preparation, label the IRE1-HaloTag protein with a Janelia Fluor 549 HaloTag ligand (Promega Cat# GA1110). Note: It is a good idea to grow up a parallel plate of unedited parental cells (containing no IRE1-HaloTag) and label them with dye in the same way so that they can serve as a control during FACS. The unedited cells will be very important in setting up gates since IRE1 is a low-abundance protein and your fluorescence signal will be very weak, so distinguishing it from the baseline will not be trivial without a proper control. 4.1.1  Dissolve the Janelia Fluor 549 HaloTag dye in anhydrous DMSO to prepare a 1 mM stock µl of the 1 mM dye stock solution into a 15 mL conical Add mL of pre-warmed growth medium to the same tube, taking care to mix well by pipetting up and down to sure the dye is This will result in a dye Aspirate the medium from the growing cells and replace it with the Place the cells in the incubator and the proceed for 1 the plate with PBS to remove Trypsinize the cells, quench the trypsin with an equal volume of fresh growth medium, and pellet the cells by gentle centrifugation (300 RCF for 3 Resuspend the pellet in FACS medium (for U-2 OS, PBS supplemented with 0.5% FBS seems to work Place the cells on the cells. There are two a plate sort, directly seeding clones into wells of a 96-well plate, or all clones together and seeding them all into a single well or As our plate was so we clones together and seeded them into a single well of a 6-well plate. The exact protocol for the plate sort will depend on your specific FACS Note is recommended to use medium FBS of 10% to of single clones Note 2: As IRE1 is a low-abundance protein with the Janelia Fluor cells a fluorescent To edited cells from we recommend using a FACS in two different (e.g. and a refer to the to see what a population like with note the low editing Place the cells in the incubator and them recover for 1-2 growth medium every 3-4 If you seeded all cells into a well of a 6-well plate or 10 cm Keep checking the clones the microscope until they use cloning to and the individual clones as in section of a If you the cells into wells of a 96-well cell growth in the wells Once a well high cell and the cells from that Once the clones have been check the and expression of edited IRE1α by Western using the Cat# to the edited IRE1 will run on a gel making the to In a good clone, for expression levels of IRE1-HaloTag that are as as to of IRE1 in the parental cells, and a of IRE1 Perform on the of the protein as needed of to the ER the & stress human IRE1α through of into and the stress K. L. CRISPR/Cas9 genome to the for in in & and of IRE1α. & A and to of

  PublisherDOI

Frequent coauthors

• Peter Walter

  University of California, San Francisco

  27 shared

• Bryan Faust

  University of California, San Francisco

  18 shared

• Nevan J. Krogan

  Gladstone Institutes

  18 shared

• Ahmet Yıldız

  University of California, Berkeley

  17 shared

• Nick Hoppe

  University of California, San Francisco

  16 shared

• Ishan Deshpande

  Google (United States)

  12 shared

• Christian B. Billesbølle

  University of California, San Francisco

  12 shared

• Smriti Sangwan

  University of California, San Francisco

  12 shared

Education

• Ph. D., Biophysics Graduate Group

  University of California, Berkeley

  2016

• B.S., Cell Biology and Molecular Genetics

  University of Maryland, College Park

  2010

Awards & honors

• John S. Swenton Award for Outstanding Teaching