Discovery Kit: dSTORM in cells

The Discovery Kit for direct Stochastic Optical Reconstruction Microscopy (dSTORM) in cells is the definitive resource for researchers looking to perform single- or dual-color dSTORM of cellular targets. It consolidates high-quality reagents with a robust starting-point protocol and guidance on how to optimize according to your targets of interest. In combination, these elements facilitate expert-standard sample preparation of almost all cellular protein targets to be imaged with dSTORM. In order to get the best results with the kit, it is important to consider the guidance on protocol selection before your first experiment and on protocol optimization during any subsequent optimization experiments. Similarly, while the secondary probes provided with the kit are validated to provide excellent high-specificity, low-background staining, it is essential that any user-provided primary probes perform to a high standard to achieve best results.

In this guide we walk you through important considerations when planning your experiment and choosing the right reagents for you. The Discovery Kit: dSTORM in cells  contains the reagents and buffers required to label a broad range of proteins and cell types. The kit is shipped at 4°C and should be stored as indicated (pages 5-6), except for the dSTORM Imaging Buffer, which is stored at -20°C.

There are two versions of the kit, briefly described below, when the exact differences between them are discussed in the next page under the “Choosing your fixative” section:

• The standard Discovery Kit (STORM DISC-M) includes Fixative A, which is paraformaldehyde (PFA).
• The Discovery Kit with Strong Fixative (STORM DISC-S) includes the option to make Fixative B, made of PFA supplemented with glutaraldehyde for stronger fixation.

An overview of the sample preparation workflow is illustrated below (Figure 1). Both the standard workflow (Fixative A) and the workflow with Strong fixative (Fixative B and Quenching) may be used with or without membrane permeabilization. In both versions the next steps after blocking include staining with primary antibodies provided by the user (up to 2 targets), and dye-conjugated secondary antibodies provided by ONI. This is followed by imaging with ONI’s two-component imaging buffer, composed of dSTORM Imaging Buffer Part A and dSTORM Imaging Buffer Part B.

Figure 1

The Discovery Kit is compatible with different formats of plates and culture dishes. It is important to use a microscopy slide with the following features:

• Has a glass imaging surface (do not use plastic surfaces).
• Has the correct thickness of glass. For the Nanoimager and Aplo Scope this is #1.5H (170 μm).
• Allows you to add and remove the dSTORM buffer, i.e. not sealed.

ONI has verified the use of Ibidi µ-Slide 18 Well Glass Bottom chambered coverslips (Cat #81817) or µ-Slide 8 Well Glass Bottom (Cat #80827).

There are two versions of the Discovery Kit with different fixatives:

1. Discovery Kit: dSTORM in cells (STORMDISC-M)
   Fixative A (4% PFA)
2. Discovery Kit: dSTORM in cells with strong fixative (STORMDISC-S)
   Fixative A (4% PFA) + 0.2% glutaraldehyde

Here we provide some general guidelines for fixative selection. We recommend that the users search the literature to see if optimized fixation conditions have been reported for their protein of interest.

Intracellular proteins
Almost all intracellular proteins can be fixed with Fixative A. We, therefore, recommend the standard Discovery Kit (STORMDISC-M) for researchers studying intracellular proteins. Cytoskeletal proteins, such as tubulin, are an exception, and are optimally fixed with Fixative B, typically using a pre-extraction step (see ‘Expanded Protocols’)

Plasma membrane proteins
For many plasma membrane proteins 4% PFA (Fixative A) is not sufficient for cross-linking and immobilization¹. Insufficient fixation of membrane proteins can lead to antibody-induced protein clustering and can impact data quality^2,3. Fixative B, which contains 4% PFA with the addition of 0.2% glutaraldehyde has been shown to preserve the native structure of many membrane proteins. However, unfortunately, not all proteins can be fixed with Fixative B as it can disrupt some epitopes and impact antibody recognition.

When optimizing the protocol for a new plasma membrane protein we recommend comparing Fixative A and Fixative B using the Discovery Kit with Strong Fixative (STORMDISC-S). If you achieve an equal or better signal using Fixative B, then proceed with Fixative B. However, if you achieve a weak signal with Fixative B but strong signal with Fixative A, proceed with the latter.

Permeabilization of the cell membrane is essential in order to stain intracellular targets. For this, a simultaneous blocking-permeabilization step is performed by supplementing the Blocking Solution with Permeabilization Concentrate. The Staining and Washing Solutions are also supplemented with Permeabilization Concentrate at a lower concentration in order to decrease non-specific binding of antibodies.

When the aim is to exclusively detect cell surface proteins, you should not permeabilize, as the detergent will damage the membrane and may change the distribution of your protein of interest. Permeabilization Concentrate should not be used at all in these experiments.

Below are examples of targets and the recommended solution usage for them:

For this kit, you need to provide your own primary antibodies based on the targets you want to study. We recommend that you perform a titration experiment between 1-20 µg/mL for any new primary antibody in order to determine the optimal concentration in your experiment. Include an isotype control to determine the background levels from non-specific binding of the antibodies or anything auto-fluorescent in the sample. This is particularly important when working with low expression proteins⁴, where the number of localizations might be close to background levels.

The ONI supplied secondary antibodies are conjugated to CFⓇ583R or AZDye™647 to enable 2-color dSTORM detection (Figure 2). These dyes exhibit optimal brightness, photostability and blinking kinetics with minimal spectral overlap, allowing two-color dSTORM imaging. Table 4 provides a list of the secondary probes offered when purchasing the Discovery Kit: dSTORM in cells.

The fluorescently labelled secondary F(ab’) 2 antibodies included in the ONI kit are compatible with the detection of primary antibodies raised in mouse, rabbit and rat hosts. The concentration of the antibodies has been validated for a broad range of targets and provides optimal labelling density and signal-to-noise ratio.

When using two different primary antibodies in one sample, they must be raised in different species as they require different secondary antibodies. These will correspond to the secondary antibody-dye pairs provided in the kit e.g., rabbit and mouse or mouse and rat, etc.

The kit has been validated for the detection of proteins in mammalian cellular compartments such as the nucleus, cytoplasm, and in organelles like mitochondria, in addition to proteins associated with the plasma membrane (examples can be found at www.oni.bio/dstormdiscoverykit). The secondary probes have been tested for the detection of mammalian protein targets in immortalized (U2OS, NIH/3T3, COS7, Jurkat) or primary (embryonic rat cortical neurons, primary human T-cells) adherent and non-adherent cells. 

*Aliquot the stock after the first use and store the aliquots at −20°C.

For simultaneous two-color dSTORM imaging select one CFⓇ583R secondary and one AZDye™647 secondary, of different species, according to your available primary antibodies. These antibodies arrive lyophilized and should be stored at -20C until resuspension, then stored at 4C for up to 3 months. See instructions in protocol for details.

• Pipettes (p10, p20, and p200)
• Pipette Tips (10, 20, and 200 μL)
• 1.5 mL microtubes
• Timer
• Water bath or heat block
• Vortex mixer
• Microcentrifuge
• Aspirator or vacuum pump (optional)
• Lens cleaning paper
• Objective oil
• Nanoimager, including magnets to hold the slide on the stage plate.

1. Primary antibodies against targets of interest raised in mouse, rabbit or rat hosts
2. Isotype-control antibodies (serve as negative control)
3. Cells cultured on high-precision coverslips or glass bottom chambers
4. Freshly prepared bead slide. Instructions can be found on the ONI Service Desk here
5. 50 μL of ultrapure water per secondary antibody

The reagent and wash volumes in the protocol are for working solutions of 100 µL per well, for example when using an Ibidi µ-slide 18-well glass-bottom slide. If using larger or smaller wells, adjust the volumes accordingly, as indicated in the table below. The listed volumes are indicative of one well. Note that only the Ibidi plates are compatible with the Nanoimager and with Aplo Scope.

• Add or remove reagents carefully using fresh pipette tips.
• Avoid touching or disturbing the cells on the surface.
• Do not allow the surface to dry out between steps.

Unless otherwise indicated, all incubations are performed at room temperature.

• Ensure the Nanoimager or Aplo Scope has been powered on and allowed to reach temperature equilibrium before proceeding.
• Instructions on setting up the Nanoimager and Aplo Scope can be found
  here, and you can also use the information provided to you by the ONI team when installing the microscope.
• Permeabilization: If staining intracellular proteins, it is essential to supplement the Wash Buffer, Blocking Solution, and Staining Solution with Permeabilization Concentrate as indicated in the stepwise protocol.
  • You can supplement your stocks when you first open the kit and keep them at 4°C for future uses, OR you can prepare your reagents each time you use the kit. Choose the latter if you are not the only kit user or if you intend to use the kit to stain targets of different cellular compartments.

As a rule of thumb, use this formula to estimate how much supplemented Wash Buffer you need:

• Set the temperature on the Nanoimager or AploScope approximately one hour before imaging to allow the system to reach a stable operating temperature.
• It should be noted that although the protocol outlined below will produce good results for most targets with little or no further optimization, some targets require additional steps or components that are not in the core protocol (e.g. pre-extraction). An overview of the most common such expansions to the core protocol is given in the ‘Expanded Protocols’ section at the end of this document. It is recommended to review this guidance before beginning work with a new target in order to assess whether such expansion is necessary.

Fixation and quenching – total time: 20 – 25 minute

Bring Wash Buffer, Blocking Solution, and Staining Solution to room temperature before starting.

The protocol specifies a working solution volume of 100 µL per well, for example when using an Ibidi µ-slide 18-well glass-bottom slide. If using larger or smaller wells, adjust the volumes accordingly, as indicated in table 5.

1. Transfer the amount of Fixative A needed for the experiment to a microtube (number of wells * volume per well + 10% excess, using Table 5).
2. Warm the fixative to 37 °C on a heating block or water bath for at least 10 minutes. Larger volumes (>1.5 mL) will require longer to warm.
3. Remove growth medium and immediately apply 100 µL pre-warmed fixative to each well.
4. Incubate for 10 minutes at room temperature.
5. Remove fixative and immediately wash with 100 µL Wash Buffer three times (no incubation needed).

Pause point: Samples can be stored for up to 1 week at 4 °C in Wash Buffer without permeabilization concentrate. Seal slides with parafilm to avoid drying out when storing.

1. Prepare Fixative B within one hour of starting the protocol according to the following instructions:
   1. Calculate the amount of fixative needed for your experiment (number of wells * volume per well + 10% excess, using Table 5. Be sure to keep the remaining Fixative A for the post fixation step).
   2. Dilute Fixation Supplement 125-fold in Fixative A in a fresh microtube. e.g., mix 496 µL Fixative A with 4 µL Fixation Supplement.
   3. After the first use, aliquot Fixation Supplement into 50 µL aliquots and store at −20 °C.

2. Warm the fixative to 37 °C on a heating block or water bath for at least 10 minutes. Larger volumes (>1.5 mL) will require more time to warm.
3. Remove growth medium and immediately apply 100 µL pre-warmed fixative.
4. Incubate for 10 minutes at room temperature.
5. During this incubation, prepare Quenching Solution:
   1. Use a fresh microtube and dilute the Quenching Concentrate 125-fold in Wash Buffer, e.g., Mix 124 µL Wash Buffer with 1 µL Quenching Concentrate. You will need 100 µL Quenching Solution for each well.

6. At the end of the incubation with Fixative B, remove fixative and immediately wash with 100 µL Wash Buffer three times (no incubation needed).
7. Apply 100 µL Quenching Solution to each well and incubate for 7 minutes.
8. Remove Quenching Solution and wash with Wash Buffer three times (no incubation needed).

Pause point: Samples can be stored for up to 1 week at 4oC in Wash Buffer. Seal slides with parafilm to avoid drying out when storing.

Blocking and staining – total time: 210 – 220 minutes

If staining intracellular proteins, add Permeabilization Concentrate to the Wash Buffer, Blocking Solution, and Staining Solution as described below. Wells with permeabilized cells will always be washed in Wash Buffer + Permeabilization Concentrate.  If preparing samples with different permeabilization requirements simultaneously, use separate wash, blocking, and staining solutions for each.

Add the Permeabilization Concentrate directly to the ONI-supplied stock bottles or to the Wash Buffer you set aside at the beginning of the assay using the following ratios:

2. Remove Wash Buffer.
3. Apply 100 µL Blocking Solution (supplemented or unsupplemented with Permeabilization Concentrate as described above).
4. Incubate for 60 minutes at room temperature.
5. During the incubation, add your primary antibodies to the Staining Solution at the desired concentration (with or without Permeabilization Concentrate). Recommended concentrations are 1–20 µg/mL, however this requires optimization for each antibody clone. If staining two targets in one sample, prepare one Staining Solution with both antibodies, each at its appropriate concentration, as you determined in your optimization process.
6. Remove the Blocking Solution and apply 100 µL of Staining Solution containing primary antibodies.
7. Incubate for 60 minutes at room temperature.
8. During this incubation, switch on the Nanoimager or the Aplo Scope to allow them enough time to warm up to a stable temperature.

1. 1. Turn on the Nanoimager or Aplo Scope
   2. Activate temperature control and set the value to 32°C
   3. Allow the microscope to heat to 32°C for at least 60 minutes
   4. Perform channel mapping for 2-color calibration if performing multi-color SMLM (instructions)  (this step should be done once the temperature is stable)

9. Wash with 100 µL Wash Buffer three times, incubating for 1–5 minutes in between each wash.
10. Dilute the secondary antibodies 100-fold in Staining Solution (with or without Permeabilization Concentrate). If staining two targets in one sample prepare one Staining Solution with both antibodies and adjust volume accordingly, e.g. 490 µL Staining Solution + 5 µL antibody A + 5 µL antibody B. Protect this solution from light until ready to use.
11. Remove Wash Buffer and apply 100 µL Staining Solution containing the secondary antibodies.
12. Incubate for 60 minutes at room temperature protected from light.
13. Wash with 100 µL Wash Buffer three times, incubating 1–5 minutes in between each wash.

Post fixation – total time: 10 – 15 minutes

1. Remove Wash Buffer.
2. Apply 100 µL Fixative A.
3. Incubate for 10 minutes at room temperature protected from light.
4. Remove Fixative A.
5. Wash with 100 µL Wash Buffer three times (no incubation needed).
6. Samples are now ready for imaging.

Pause point: If you cannot proceed to imaging right away, samples can be stored in Wash Buffer for up to 1 week at 4°C protected from light. Seal slides with parafilm to avoid drying out when storing.

This section provides guidance on the key steps to acquire and analyze a dSTORM image on the Nanoimager. If you haven’t used the Nanoimager before, get in touch for some basic training. Ideally, you have already switched the Nanoimager or Aplo Scope on, and their temperature has been equilibrated. If so, follow the steps in “dSTORM imaging” section to set a light program. If not, do it now, and allow at least 60 minutes for the process.

1. Turn on the Nanoimager
2. Activate temperature control and set the value to 32oC
3. Allow the microscope to heat to 32oC for at least 60 min
4. Perform channel mapping for 2-color calibration (instructions)

Before you prepare the Imaging Buffer and apply it to the cells, make sure you know how to use the microscope, as the dSTORM Imaging Buffer is sensitive and is only stable for about 1 hour. Remember, you can store your stained samples for up to 1 week at 4°C if needed.

1. Thaw one vial of dSTORM Imaging Buffer Part A at room temperature for 10 minutes.
2. Spin the tube down using a tabletop centrifuge for 10 seconds.
3. Carefully mix the solution with a pipette, making sure not to introduce bubbles.

4. Add 99 µL dSTORM Imaging Buffer Part A to a new microtube for the well that is to be imaged. This should be allowed to reach room temperature or, preferably, the imaging temperature of the microscope before continuing. This will minimize thermal drift during imaging.

5. Remove dSTORM Imaging Buffer Part B from -20°C. Ensure you re-seal the plastic bag from which it was removed. If this is the first time using this Part B Buffer, resuspend the buffer following steps 6-8. If the Part B is already resuspended, skip to step 9. 
6. Add 25 µL of Part B Resuspension Buffer to the vial. Slowly add a single drop to the center of the pellet, and apply the rest to the sides of the vial to ensure complete dissolution of the pellet.
7. Put the rehydrated vial on ice for 5 minutes.
8. Gently mix the re-hydrated pellet by pipetting up and down. The vial can be removed from the ice bucket for this process. Do not introduce bubbles or air. Do not shake or vortex the vial.
9. Take 1 µL of the resuspended dSTORM Imaging Buffer Part B and add it to the 99 µL of dSTORM Imaging Buffer Part A in the fresh microtube. Once the required volume is removed, put the remaining resuspended dSTORM Imaging Buffer Part B at -20°C and use within 3 months.
10. Remove Wash Buffer from wells.
11. Apply 100 µL of dSTORM Imaging Buffer mix to the well you are about to acquire. Pipette gently up and down to generate a homogenous solution before adding to the imaging well. Note that we recommend adding dSTORM Imaging Buffer mix and imaging one well at a time.
12. Cover chambered slide with lid to minimize evaporation and exposure to oxygen.
13. When preparing dSTORM Imaging Buffer sequentially, dSTORM Imaging Buffer Part A can be kept on ice during this time. dSTORM Imaging Buffer Part B should always be kept at -20°C and only taken out when needed.
14. Proceed to imaging your sample.

1. Mount the sample onto the microscope stage, find focus and set Z-lock (instructions)
2. Set a light program for one secondary antibody or a simultaneous light program for two secondary antibodies (instructions)

Below are some general Nanoimager and Aplo Scope imaging starting conditions, but you will need to find the right settings for your experiment:

• Exposure time – 30 ms
• Number of steps – 1
• 640 laser – minimum 50 mW
• 561 laser – minimum 50 mW
• Number of frames – 10,000–20,000 (depending on sample density)

3. Press “Save and Activate” to store the light program.
4. Set the file and folder name.
5. Set the number of repetitions of the light program to 1.
6. Find your desired field of view (FOV) using low intensity illumination.
7. Find the optimal focal plane and illumination angle for the structures of interest.
8. Press “Acquire” when ready to start.

Exposure time
30 ms is a good starting point for most targets as it is sufficiently long to allow robust photon-detection from individual molecules, but sufficiently short to allow separate blinking events to be localized independently.

In cases where the imaging target is extremely dense, it may be necessary to reduce the exposure time in order to reduce the frequency at which multiple nearby molecules are localized as one in the same frame (due to overlapping blinking events). In such cases, exposure time can be reduced to 20 or 10 ms, with a corresponding adjustment in laser power (see below).

It is typically not necessary to use longer exposure times if performing of dSTORM CFⓇ583R- or AZDye™647-labelled targets, however if an alternative imaging methodology is being used that has slower blinking kinetics (such as DNA-PAINT), then increasing exposure time to 50 or 100 ms may improve imaging outcomes.

Laser power
In most cases 50 mW is sufficient to produce robust blinking for both CFⓇ583R- or AZDye™647-labelled targets. However if using a very stringent illumination angle in order to restrict illumination only to highly proximal parts of the sample it may be necessary to increase laser power to account for the reduction in at-sample photon density caused by high illumination angles. To optimize this, simply increase laser power in continuous illumination mode until robust blinking is observed, record the laser power at which this is achieved, and then input this value for the relevant laser in the light program.  

Similarly, if exposure time is reduced below 30 ms it is recommended to increase laser power to ensure that sufficient light is collected from each molecule within the shorter frame. A simple starting point is to increase power by an equivalent proportion to the decrease in exposure time: e.g. if halving exposure time to 15 ms, then double increase laser power to 100 mW etc. This will also have the effect of ensuring that the blinking rate remains high so that most blinking events will be restricted to a single or small number of frames.

Frame number
The optimal number of frames to collect depends largely on the abundance of the target imaged, and the acceptable amount of time available for acquisition. Broadly, samples with more abundant target(s) should be imaged for a greater number of frames than those with less abundant target(s), as this ensures a comprehensive sampling of the molecules therein. For example, highly abundant, dense targets such as Tubulin, Nuclear Pore Complexes, Actin etc. should be imaged for 20k frames or more to achieve best results, whereas lower abundance targets such as surface receptors like HER2, CD19, or TCR may be comprehensively imaged in 5-10k frames. 

As a general rule, if the amount of blinking ongoing at the end of a light program is very little (i.e. localizations/frame reduced by >95% compared to localizations/frame after 500 frames. Localizations/frame value is visible during acquisition in NimOS) then the number of frames acquired may be reduced, whereas if it is very substantial then the number of frames may be increased. If performing single-color or simultaneous two-color imaging then there is no drawback to increasing frame number aside from the additional time it takes to perform one acquisition, however whether this is necessary will depend on the experiment and the desired information needed from the end data.

If adjusting the exposure time it is generally recommended to adjust the total frame number to keep the overall imaging time constant – i.e. if halving from 30 ms to 15 ms exposure, then also double the number of acquired frames. This ensures that the total information collected is not reduced due to potential increases in single blinking events crossing multiple frames.

Image analysis should be performed on ONI’s cloud-based platform, CODI, where users can upload, organize, filter, and analyse their data. This section provides guidance on the key steps in CODI. For more information, please refer to the How to Guide on our CODI Help section.

1. Upload the acquired datasets using the CODI Desktop Uploader. More information on how to download the CODI Desktop Uploader and how to use it can be found here.
2. Once the upload has finished, go to CODI and navigate to your dataset.

3. Click the “Add a new analysis” tab, select the Clustering App and the Discovery Kit: dSTORM settings and click run all steps. This runs the following workflow:

1. 1. Drift Correction
   2. Single molecule filters
      1. Frame index: >300
      2. Photon count: >300
      3. Sigma value: 50-250
      4. Localization precision: 2-25
   3. Regions of Interest (ROI) (optional)
   4. Clustering
      1. DBSCAN
      2. Apply filter of clusters to restrict analysis to desired cluster morphology

These settings should serve as a starting point for your analysis, and it is likely that you will need to adjust the localization filters and the clustering settings. If you would like to use the counting tool, select the Clustering and Counting App and the corresponding Discovery Kit: dSTORM settings. To learn more about drift correction, localization filtering and clustering please see our CODI how-to articles.

• Increase the primary antibody concentration.
• Increase the duration and decrease the temperature of primary antibody incubation (i.e., 4°C, overnight).
• Change the fixative from Fixative A to Fixative B, or vice versa.
• Confirm that sample pre-extraction (see ‘Expanded Protocols’) is not required for your experiment and adjust accordingly if it is.

• Reduce primary antibody concentration.
• Increase the duration and decrease the temperature of primary antibody incubation (i.e., 4°C, overnight). This should be performed in conjunction with decreased primary antibody concentration.
• Increase the number and duration of the washes after primary antibody labelling.
• Test alternative primary antibodies.
• Confirm that supplemental blocking considerations (see ‘Expanded Protocols’) are not relevant for your experiment and adjust accordingly if they are.
• When it is difficult to differentiate between signal and background, use an appropriate isotype control antibody to report pure non-specific staining.

• If the stained sample is bleaching rapidly, consider doubling the concentration of dSTORM Imaging Buffer Part B in the dSTORM Imaging Buffer mix.
• If the rate of blinking is slow, confirm that laser power is sufficient to achieve robust blinking given your chosen illumination angle and focal plane. As a general rule, blinking is of good quality if the majority of blinking events only persist for a small number of frames (1-5) and if the vast majority of fluorophores are in a blinking regime. If you are still able to identify the underlying structure by eye this suggests that too many fluorophores are in a non-blinking regime. Note that it may take several hundred frames for the full population to begin blinking properly.

The protocol outlined above is applicable for a majority of targets, however in some cases it should act more as a starting point from which a fully target-optimized protocol may be developed. Optimization in most cases will consist of adjustments to the steps described above – e.g. optimizing primary antibody concentration, staining time, staining temperature etc. – however in a minority of situations additional steps and/or supplementary reagents may be required to fully optimize to a given target. Three of the most common expansions to the core Discovery Kit protocol are outlined below.

Pre-extraction is an optional step that can be used to remove a proportion of the soluble cytosolic molecules within a cell prior to fixation. This is typically used either 1) to remove the cytosolic fraction of a target but leave behind a non-cytosolic fraction of interest (e.g. removing soluble tubulin monomers to improve imaging of microtubules); or 2) to reduce the density of the fixed cytosol in order to improve probe accessibility to difficult-to-reach targets, such as those in the nucleus or other dense subcellular compartments. Pre-extraction should never be used with soluble cytosolic targets, or with targets that are dependent on membrane interactions for their organization (e.g. endosome-associated proteins etc.).

Whether Pre-Extraction Solution should be supplemented with Fixation Supplement depends on the sensitivity of the intended target to pre-extraction. For cytoskeletal proteins, such as tubulin, pre-extraction without fixative often leads to destabilization of cytoskeletal structures and hence a deterioration in staining fidelity, and so it is recommended to include fixative in such cases. For difficult-to-reach targets, such as nuclear pore components, pre-extraction can be performed without fixative as they are typically not sensitive to disruption by detergent during a short pre-extraction step.

The precise concentrations of Permeabilization Supplement and/or Fixation Supplement can be further optimized according to cell type and target as needed, using the above values as a starting point.

Pre-extraction can be performed as follows:

1. Prepare the Pre-Extraction Solution:
   1. Dilute the Permeabilization Concentrate 50-fold in Wash Buffer.
   2. Optional Step: Add Fixation Supplement in a 250-fold dilution according to whether simultaneous fixation and pre-extraction are required (see guidance below).
2. Remove growth medium and immediately add 100 µL Pre-Extraction Solution.
3. Incubate for 15-45 seconds at room temperature.

Remove Pre-Extraction Solution and immediately replace with 100 µL of relevant fixative. Continue to follow the normal fixation protocol and downstream steps thereafter.

The Discovery Kit Blocking Solution efficiently prevents most non-specific protein deposition within the sample. This ensures low background when using protein-based probes. In cases where nucleic acid probes are used, such as DNA hybridisation probes, DNA-PAINT strands, or nucleic acid aptamers, it may be necessary to supplement the Blocking Solution with a nucleic acid-specific blocking agent. We recommend a reagent such as Salmon Sperm DNA used at ~5% w/v added directly into the Blocking Solution and otherwise used as described. Such reagents are not included within the Discovery Kit and must be supplied by the user.

In some cases, it may be necessary to block specific protein-protein interactions, which will not be blocked by the Discovery Kit Blocking Solution. This is typically required in cases where a non-target molecule within the sample exhibits specific affinity for the probe(s) used. The most common example of this is in specific Fc Receptor (FcR)-antibody interactions if working with FcR-bearing cells of the same species as the primary antibodies. In such cases, the Blocking and Staining Solutions should be supplemented with an appropriate agent that will block the undesired interaction, e.g. commercially-available FcR-neutralizing antibodies. Such reagents are not included within the Discovery Kit and must be supplied by the user.

In rare cases, targets may require a specific base buffer to optimally stabilize them during fixation. This is generally only the case when pre-extraction is used and hence normal cellular conditions are disrupted before the sample is robustly fixed. For example, for optimal fixation of actin or actin-associated molecules it is often recommended to use PEM (PIPES, EGTA, MgCl2) buffer as the base solution for pre-extraction and fixation. In cases where this is preferable, simply use the relevant base buffer in place of the Discovery Kit Wash Buffer in the Pre-Extraction Solution, and in place of Fixative A in the Fixation Solution. Following fixation, the normal Discovery Kit Wash Buffer may be used as usual.

Fully optimized protocols for a number of common targets, including tubulin, actin, and TOMM20, can be found on the ONI Resource Library: https://oni.bio/resources/resource-library

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4. Nerreter T, Letschert S, Götz R, Doose S, Danhof S, Einsele H, Sauer M, Hudecek M. Super-resolution microscopy reveals ultra-low CD19 expression on myeloma cells that triggers elimination by CD19 CAR-T. Nat Commun. 2019; 10(1): 3137. doi: 10.1038/s41467-019-10948-w