FAQ: Assay/Workflow

No, the Microscoop Mint platform requires fixed cells. Fixation ensures that proteins of interest remain in stable locations during the photolabeling process, enabling consistent protein composition within Regions of Interest (ROIs) across all fields of view (FOVs).

The photolabeling process would induce unwanted responses by live cells, altering results as photolabeling progresses. Tissue sections can be fresh-frozen or formalin-fixed, parafin-embedded (FFPE).

We have not yet validated the Microscoop assay in plant cells or tissues. However, our reagents are species-agnostic, and successful application would depend on two key factors:

The ability to achieve high-quality immunofluorescence staining for accurate ROI masking.

Effective membrane permeabilization to allow the photolabeling reagent to penetrate plant cell structures.

We recommend conducting preliminary feasibility studies and welcome inquiries for collaborative validation. Please contact support@syncell.com for guidance.

Our workflow has been validated for human and mouse samples. Our chemistry is species agnostic. Please be reminded to prepare your own database (protein sequences as a FASTA formatted file) for MS data analysis

No, the Microscoop workflow is designed for bottom-up proteomics. We perform protein digestion and purification prior to LC-MS/MS, meaning intact proteins are not analyzed in their full-length form.

If the post-translational modification (PTM) of interest has a high occupancy (e.g. > 20%), the Microscoop could be well-suited for these experiments. However, for low-occupancy PTMs, it becomes challenging to identify and quantify them due to their low abundance. In such cases, an additional enrich step is required of photo-biotinylated proteins to enrich the specific PTMs, making the workflow more complex.

Our current reagents support photobiotinylation and collection of protein only.

No, Microscoop is designed to pool protein from many sites to create a representative proteome of similar structures. This requires collecting enough protein for mass spectrometry analysis.

Yes. Primary cells or cell lines may be used as long as they are fixed and adherent to the glass slide.

Yes. Endogenously expressed fluorescent proteins may be used if they are fixed and permeabilized.

For each photolabeled experimental sample, an unlabeled control sample should be run in parallel. This control does not undergo photolabeling and is used to account for non-specific biotinylation or background binding to streptavidin beads.

Data from the unlabeled control is essential during analysis to identify significantly enriched proteins in the photolabeled samples by comparison, ensuring high confidence in the specificity of detected proteins.

We use nuclear staining and the nuclear proteome as a reference standard to evaluate the specificity and success of the assay. This positive control allows users to confirm effective photolabeling and proper workflow execution.

Please refer to the Synlight-Rich IFU for detailed instructions on performing this analysis and interpreting results.

Our workflow is compatible guanidine-based homogenization as long as the concentration of guanidine HCl is below 1M before performing the pull down assay.

You can achieve this by co-staining the two cell types with distinct fluorescent markers. The overlapping signal from both markers is used to generate a mask that identifies the contact surface between the two cell types.

Only this overlap region will be targeted and photolabeled, enabling specific collection of proteins from cell–cell contact sites.

You can assess XY labeling precision through a simple photobleaching test—monitoring the precise area where photobleaching occurs to evaluate targeting accuracy. We usually use Nucleus or Nucleolus staining in fixed cells as the target to evaluate precision.

For Z-axis labeling precision, perform an efficiency test using nucleus in fixed cells as target and analyze the biotinylation efficiency (intensity) using ImageJ software or equivalent image analysis tools. This allows you to quantify labeling specificity and depth accuracy without requiring a confocal microscope.

Using nuclei-labeled cells as an example, more than two times of experiments (biological replicates) can increase the enrichement from ~84% to 99%.

Fixed cells or tissue sections can be used. We have validated our assays and reagents on fresh-frozen sections up to 20 μm thick and formalin-fixed, parafin-embedded (FFPE) sections up to 10 μm thick.

Using the Synlight-Rich kit with Microscoop Mint, lateral (XY) labeling resolution is approximately -350 nm and axial (Z) resolution is approximately ~1500 nm.

We require 50 - 400 ug total protein input before using the Synpull kit to perform enrichment and purification. The amount of input protein for streptavidin pull down before LC-MS/MS depends on the size and abundance of your target region of interest. For example, if you are studying an ROI as large as a nucleus, which is present in every cell, ~50 ug of input protein will be sufficient. For smaller condensates (e.g. stress granules, early endosomes), we suggest ~ 200 ug to up to ~ 400 ug protein for quality data.

To determine how many slides you will need for your experiments, please contact support@syncell.com. A field application scientist will organize a meeting with you to discuss experimental planning and determine the number of slides required per project.

This feature is not available at this time.

We recommend FFPE samples are prepared as 5-10 um sections and fresh-frozen tissues are 10-20 um in thickness. Using tissues greater than 20 um in thickness are beyond the focal capabilities of the system, resulting in imprecise photolabeling.

We typically target a photolabeling pixel count exceeding 1×10^8 and aim to harvest at least 100 µg of total protein. This yield supports the recovery of 100–150 ng of protein for downstream mass spectrometry (MS) analysis.

From experience, seeding cells to ~80% confluency in a one-well chambered coverslip typically yields 80–100 µg of protein, though this can vary by cell type.

The Synpull workflow includes multiple washing and desalting steps to ensure sample purity, which can result in protein loss. To compensate for this and ensure sufficient recovery for mass spectrometry, we recommend starting with at least 100 µg of protein prior to the pull-down.

This starting amount ensures you obtain enough material—typically 100–150 ng of enriched protein—for successful MS analysis.

We typically start with ~100 µg of total protein for immunoprecipitation, resulting in approximately < 200 ng of peptide digest at the end of the workflow—sufficient for LC-MS/MS analysis.

In some cases, lower amounts may be adequate, depending on the sensitivity and type of mass spectrometer used. Please consult with your MS facility to determine the minimum input required for your specific instrument and analysis needs.

We aim for approximately 100 ug of total protein before using the Synpull kit, and recommend 200 - 400 ug for new projects that have yet to be tested experimentally. However, we also need to consider the ROI mass necessary to obtain meaningful data. We typically determine this by using pixel counts values calculated by the Microscoop software. We recommend a range of 10^7 to 10^8 pixels be captured to obtain enough ROI mass for data analysis. Please note that these estimations may vary depending on the sensitivity of the mass spectrometer used.

We can detect proteins in low abundance with as few as 100 copies per cell.

Sample preparation time can vary depending on your experimental design. All samples must be fixed and either immunofluorescently stained or endogenously express fluorescent proteins to be compatible with Microscoop Mint photolabeling.

Typical preparation—including fixation, permeabilization, staining, and probe incubation—can take several hours to overnight, depending on protocol specifics. For detailed timelines and step-by-step instructions, please refer to the sample preparation section in the Synlight user guide.

Yes, you may use a different imaging channel for verification, as long as the fluorophore is spectrally distinct from the photolabeling channel used in your experiment. This ensures clear signal separation and accurate imaging results.
For guidance on compatible fluorophores and channel selection, please contact your Field Application Scientist (FAS) or email support@syncell.com.

Using the Cy5 channel helps avoid unintentional photolabeling across the entire field of view during imaging. Cy5’s excitation wavelength is the farthest from the UV activation range of the Synlight-Rich reagent.

The 780 nm laser used for photolabeling only activates the reagent through two-photon excitation, which is not triggered by the LED light source used for imaging. We have validated that FITC and TRITC channels can also be used safely without affecting photolabeling.

S/N ratio > 8 is a criterion we use and recommend to confirm whether the labeling conditions are optimized for successful biotinylation. Its use is limited only to the biotinylation quality; it is not an indicator of the MS quality. For the quality of the MS result, we will usually check the liquid chromatography (LC) chromatogram and the labeled v.s. unlabeled via volcano plot of the MS result. A random distributed chromatogram suggests unbiased analysis by the spectrometer, while a right-skewed volcano plot suggests enrichment of biotinylated proteins in the labeled group as IFU 7.5 step 9 shown.

No, we do not provide a predefined list. You may use any antibodies that yield high-quality immunofluorescence staining suitable for identifying your regions of interest (ROIs). Antibody choice should be guided by your specific target proteins and sample type.

The number of kits needed is dependent on the scope of the project and may vary. Our kits contain 6 reactions, which are sufficient for 3 x unlabeled controls and 3 x photolabled samples. Please contact support@syncell.com for additional assistance.

Please note the storage conditions indicated on each box. One of the sub-components, 'K' from the Synpull kit, must be stored at -20 °C. All other items must be stored at +4 °C until use. Importantly, ensure that the streptavidin beads do not freeze at any time.

The exact composition of our reagents is proprietary. For help optimizing or troubleshooting your protocols, please contact us at support@syncell.com.

Although it is possible to prevent hydrolysis by Trypsin digestion on modified arginine (R). The production of multiple hydrolyzed peptides based on a single protein is still sufficient for identification.

The UL control is used to detect non-specific biotinylation of proteins. It uses photolabeling reagent, and follows the photolabeled (PL) sample through all steps except photolabeling by the laser of Microscoop Mint. The UL control is further used downstream of our workflow to serve as a control in our analyses.

The Synlight-Rich +Positive Control Kit provides the specific reagents needed to introduce a positive control into your Microscoop workflow. This control is useful for confirming that the assay has been successfully executed, and it can also support workflow optimization and troubleshooting.

No. We currently only support biotin-based photolabeling and subsequent streptavidin pull-down for protein enrichment and analysis.

If the PL and UL undergo the same round of mass spectrometry preparation, this is OK. But if preparation is done at different times for different PL samples, each needs its own UL control

No, the photolabeling reagent is not inherently membrane-permeable. Therefore, fixed cells must be permeabilized prior to the photolabeling step to allow the reagent to enter the cells.

Failure to permeabilize the cells can result in reduced or incomplete biotinylation of the targeted Regions of Interest (ROIs), compromising experimental results.

Using nuclei-labeled cells as an example, We estimate biotinylation efficiency is appoximately 0.1~1%.

The photoactivatable biotin reagent inserts into the alpha carbon (Cα) of amino acid residues in a non-specific manner. However, certain residues are preferentially biotinylated, including methionine, glycine, phenylalanine, and arginine. These residues tend to be labeled more frequently due to their chemical properties and accessibility during the reaction.

Syncell is currently the only source of our reagent kits - we do not have authorized dealers. We cannot offer support for customer-developed reagents or workflows.

Please contact your FAS, Territory Sales Executive or support@syncell.com for guidance.

At this time, we do not sell sub-components of our kits separately.

Please contact your local Territory Sales Executive or info@syncell.com for further information.

Photolabeling efficiency is measured by calculating the signal-to-noise ratio (S/N ratio). This involves comparing the fluorescence intensity of the labeled target region to the background signal or noise from the rest of the cell.

A higher S/N ratio indicates more specific and efficient labeling, which reflects effective targeting and minimal non-specific signal.

Photolabeled samples can be stored for up to one month at +4 °C. To maintain sample integrity during this period, we recommend adding sufficient PBS containing 0.05% (w/v) sodium azide to prevent cell dehydration and microbial contamination, while also protecting samples from light to preserve photolabeling stability.

While alternative methods that efficiently and accurately pull down biotinylated proteins may technically be used, we strongly recommend using the Synpull kit. The Synpull kit is validated for compatibility with Microscoop-generated samples, and using other pull-down methods will result in our being unable to provide technical support for your assay or data interpretation.

Reserve a small portion of the photolabeled sample to verify biotinylation efficiency using P (Verify), as outlined in the Synlight-Rich Kit IFU.

Option 1: Dot Blot Assay

Spot 1–2 µL of protein-bound beads (and biotin standards) onto a PVDF membrane.

Activate membrane with 100% methanol, soak in PBS-T for 2 min, and allow to dry.

Rinse with 0.1% Triton X-100/PBS.

Block with 5% BSA for 30 min at room temperature, wash again with 0.1% Triton X-100/PBS.

Incubate with streptavidin-HRP (1:1000) in 5% BSA at 4°C overnight.

Wash with 0.1% Triton X-100/PBS and develop using ECL substrate. Image with iBright FL1500 (Cytiva).

Option 2: LC-MS Analysis
Calculate the ratio of enriched proteins to total proteins from the LC-MS results to estimate biotinylation efficiency.

The positive control (PC) is a nucleus-labeled sample, and nuclei have a larger area/volume compared to other organelles or regions of interest, resulting in higher biotinylated protein yield per area post photolabelling.

In contrast, customer samples targeting smaller regions may require more starting protein amounts to ensure sufficient yield. Additionally, the protein-to-bead ratio (µg/µL) and incubation time (e.g., 1 hour) may need to be optimized for specific applications. For initial testing, we recommend 200–400 µg of protein with 10 µL of beads.

Biotinylation adds a monoisotopic mass of 626.2774 Da to the modified peptide. This value should be used in mass spectrometry searches, not the average mass.
However, due to the on-bead digestion workflow used in the Synpull kit, most biotinylated peptides remain attached to the beads and are not eluted. As a result, even if this mass shift is included in your search parameters, biotinylated peptides are rarely detected or appear at very low abundance.

The ultrasonic water bath is used to disperse magnetic beads and remove excess detergent. Due to the small sample volume, simple pipetting may not effectively separate beads, especially as detergent is reduced. Ultrasonication ensures the beads remain evenly suspended and prevents clumping, which is essential for efficient downstream processing.

At this time lysates produced by the Synpull kit can only be measured using the Pierce 660 nm Protein Assay (IDCR method). If other methods are capable of detecting 660 nm absorbance readings, protein concentrations may also be determined by creating a standard curve in that way as well. Please contact your FAS or support@syncell.com for further guidance.

This approach is sufficient for cell culture samples, as indirect sonication can provide enough energy for cell lysis. However, for tissue samples, a suitable sonication probe is required to ensure proper homogenization and efficient protein extraction.

Based on our experience, 30% amplitude using the Qsonica 125 model is sufficient to lyse tissue samples. Using a higher amplitude may damage the samples, so optimization is recommended if a different sonicator or power setting is used.dn optimization if other sonicating power is desired.

Incubation can range from 12-20 hours without loss of quality.

The purpose of the 2-hour digest is to reduce the amount of Streptavidin (SA) peptides in the Synpull product. After testing various enzyme concentrations and on-bead digestion times, we found that a 2-hour on-bead digestion is an optimal balance. It minimizes SA peptide amount without significantly affecting the number of protein IDs. On the other hand, we are currently developing Synpull Kit v2.0, which supports overnight (16-hour) on-bead digestion and with very few SA peptides in the pulldown product.

We have verified that the beads can be stored in the wash buffer (component E: Dilute) at 2-8°C for up to one year. For friendly operation and one-step washing the beads in bulk ensure proper bead concentration as spection (80 uL for first time using), as long-term storage result in volatilization if beads are stored without this procedure.

It is critical to maintain the lysate volume at 100 µL (e.g, from 720 µL/6 x Single-wells) to ensure the correct 100:60 volume ratio of Lysate to Lysis Buffer. This ratio ensures the proper concentration of detergent needed for effective protein extraction.

Deviating from this ratio may alter detergent levels, leading to inefficient protein extraction and protein loss during the workflow.

A 5-fold dilution is essential to provide optimal conditions for biotin–streptavidin binding. This dilution does not result in protein loss—the total protein remains in the same tube; only the concentration is reduced.

We have validated that protein identification remains consistent across concentrations of 0.125–0.625 mg/mL (e.g., 100 µg in 160–800 µL), assuming the same buffer composition. This ensures efficient and reproducible pull down performance.

We combine activation and binding into one step. It is crucial to ensure the sample pH is below 4 and perform adequate aspirate–dispense cycles (e.g., 30 cycles).

Dried peptide digests can be stored at –20 °C for up to 1 month prior to LC-MS/MS analysis.

To ensure optimal peptide stability, avoid repeated freeze–thaw cycles and store samples in a low-humidity, contamination-free environment.

Protein lysates can be stored:

At +4 °C for up to 24 hours (short-term storage), or

At –20 °C for up to 1 month before proceeding with immunoprecipitation.

For optimal results, minimize freeze–thaw cycles and process lysates as soon as possible.

Currently, the lysate produced by the Synpull Kit can only be measured using the Pierce 660 nm Protein Assay (IDCR method). If a Nanodrop capable of detecting 660 nm absorbance is available, protein concentrations can also be determined by creating a standard curve.

We would feel confident with 1.5-fold difference (Laber Free Quantification by MS results). You can put serial tissue sections on a single slide and do both at the same time to limit batch effects. The dynamic range depent on LC-MS/MS level about 3~4 order.

Lysis buffer can be changed by the customer, however SDS concentration should be less than 0.5% during AP (Affinity purification).

No. The UL (unlabeled) control reflects the natural background distribution of proteins in your sample. Final results are determined by comparing photolabeled samples to their matched UL controls, allowing us to accurately identify significantly enriched proteins.

A high percentage of protein IDs in UL samples is normal and expected, and does not impact the validity of the final analysis.

No. We do not use a cleavable biotin at this time.

We use a concentrated liquid protease inhibitor that is pre-added to Reagent Scrape (A): Scrape of the Synpull Kit. This inhibitor has been validated for long-term stability when stored at 2–8 °C, ensuring consistent protection against proteolysis without the need for fresh addition prior to use.

No. During the protein extraction and digestion process, RNA molecules are typically cleaved and degraded. As a result, only protein-derived peptide fragments are recovered and detected by LC-MS/MS. RNA is not co-recovered or analyzed in the Synpull workflow.

The Synpull protocol used for cell samples is also applicable to FFPE tissue sections. The only modification is during the lysis step: a sonication probe should be used to homogenize tissue instead of a standard sonication bath.

This ensures effective protein extraction from FFPE samples while maintaining compatibility with the rest of the SynPULL workflow.

Protein LoBind tubes are strongly recommended. After washing Streptavidin beads with Buffer J, the beads become very sticky and may adhere to standard tube walls, making it difficult to collect and resuspend them for on-bead digestion.

For larger structures (e.g., nuclei), this may have minimal impact on protein identification. However, for small-scale or low-abundance targets—such as stress granules or primary cilia—using normal tubes can result in a significant reduction in protein IDs