Discovery Kit: dSTORM in cells 2

The Discovery Kit for direct Stochastic Optical Reconstruction Microscopy (dSTORM) in cells is the ultimate kit for researchers looking to prepare samples for single- or dual-color dSTORM super-resolution imaging of cellular targets. The kit contains the reagents and buffers required to label a broad range of proteins and cell types. These high-quality reagents are combined with a robust protocol to optimize labeling of targets of interest for dSTORM imaging, facilitating expert-standard sample preparation of nearly all cellular protein targets. 

In this guide, we walk you through important considerations when planning your experiment and choosing the right reagents. To get optimal results with the kit, it is important to consider the guidance provided on protocol selection ahead of the initial experiment and on protocol optimization for any subsequent optimization experiments. Similarly, while the secondary antibodies provided with the kit are validated for excellent high-specificity and low-background staining, any user-provided primary probes must be validated for high performance to achieve the best results. 

Important: 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:

There are two versions of the kit, briefly described below: 

• 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 Strong Fixative, which is made of Fixative A and Fixation Supplement (0.2% glutaraldehyde) for stronger fixation together with a Quenching Concentrate.  

The differences between these are discussed under the “Choosing your fixative” section.  

An overview of the sample preparation workflow is illustrated below (Figure 1). Both the standard workflow (Fixative A) and the workflow with Strong Fixative 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

For simultaneous two-color dSTORM imaging, select one Cy3B secondary and one AZDye™647 secondary, of different species, according to the available primary antibodies.

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

• 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/Aplo Scope, 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 Discovery Kit is compatible with different plates and culture dish formats, with the following criteria:  

• A glass imaging surface (do not use plastic surfaces)
• The correct glass thickness. For both the Nanoimager and Aplo Scope this is #1.5H (170 μm)
• Allow the addition and removal of 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).

The composition of the fixative should be considered, knowing that certain structures or epitopes require a stronger fixation to immobilize or preserve ultrastructure. Here, we provide some general guidelines for fixative selection. However, we strongly recommend that the users search the literature to see if optimized fixation conditions have been reported for their protein of interest.

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

1. Fixative A (4% PFA) in the dSTORM Discovery Kit (SKU: STORMDISC-M) 
2. Strong Fixative (Fixative A + 0.2% glutaraldehyde) in the dSTORM Discovery kit with Strong Fixative (SKU: STORMDISC-S)

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 kit (STORMDISC-M) for researchers studying intracellular proteins. Cytoskeletal proteins, such as tubulin, are an exception and are optimally fixed with the Strong Fixative, 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 quality2,3. Strong Fixative, has been shown to preserve the native structure of many membrane proteins. However, not all proteins can be fixed with Strong Fixative 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 Strong Fixative using the Discovery Kit with Strong Fixative. If you obtain an equal or stronger signal with Strong Fixative, proceed with that condition. However, if Strong Fixative yields a weak signal while Fixative A produces a strong signal, continue with Fixative A instead. 

We recommend reading our Technical guide: Sample preparation & imaging with the ONI Discovery Kit for dSTORM in cells for further information (for Nanoimager only). 

Permeabilization of the cell membrane is essential 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 to decrease non-specific binding of antibodies.

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 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 Cy3B 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 3 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. 

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.

• Follow the instructions on setting up the Nanoimager and Aplo Scope found in the Appendix.   

• 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. 
  Stocks can be supplemented when first opening the kit and kept at 4°C for future uses, OR reagents can be prepared each time the kit is used. Choose the latter when intending to use the kit to stain targets of different cellular compartments. If quenching is necessary, i.e., if using Strong Fixative, Wash Buffer without Permeabilization Concentrate should be reserved. 

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

• Set the temperature on the microscope to allow the system to reach a stable operating temperature and avoid stage drift: 
  • Nanoimager: set the temperature one hour before imaging 
  • Aplo Scope: set the temperature two hours before imaging

While 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 expansions to the core protocol is given in the ‘Expanded Protocols’ section. It is recommended to review this guidance before beginning work with a new target to assess whether any expansion is necessary. 

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 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. 

Fixation and quenching – total time: 20 – 25 minute

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

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 Strong Fixative 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 Strong Fixative, 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 32oC: at least 1 hour for the Nanoimager, and at least 2 hours for Aplo Scope 
   4. Perform channel mapping for 2-color calibration if performing multi-color SMLM. Read the instructions: for the Nanoimager or for Aplo Scope (“Control Software – channel mapping” section). 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. If this is the first use of the kit, resuspend the provided secondary antibodies in 50 uL of ultrapure water by slowly adding a single drop to the center of the pellet, and applying the rest to the sides of the vial to ensure complete dissolution of the pellet. After allowing the pellet to rehydrate for 5 minutes, gently mix by pipetting up and down. After resuspension, store the remainder at 4C and use within 3 months. 
11. 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.
12. Remove Wash Buffer and apply 100 µL Staining Solution containing the secondary antibodies.
13. Incubate for 60 minutes at room temperature protected from light.
14. 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 or Aplo Scope. If you haven’t used our microscope before, get in touch for some basic training. Ideally, you have already switched the microscope on, and its 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.

Prior or during the incubation step for the Staining solution  

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: at least 1 hour for the Nanoimager, and at least 2 hours for Aplo Scope
4. Perform channel mapping for 2-color calibration if performing multi-color SMLM. Read the instructions: for the Nanoimager or for Aplo Scope (“Control Software – channel mapping” section). This step should be done once the temperature is stable.

Before preparing the Imaging Buffer and applying it to the cells, ensure everything is ready for imaging, as the dSTORM Imaging Buffer is sensitive and only stable for about 1 hour. Note: stained samples can be stored 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.
   Note: Never flick or vortex the dSTORM Imaging Buffer. Introducing oxygen will affect its performance
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.
   Note: We recommend imaging one well at a time.  
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. 

With a Nanoimager

1. Mount the sample onto the microscope stage, find focus and set Z-lock (instructions) 
2. Set a light program (instructions) 

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

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.

With an Aplo Scope

1. Mount the sample onto the microscope stage, find focus and set Z-lock > instructions
2. Set a multi-acquisition sequence (instructions under “Control Software – Multi-Acquisition Sequence” section) 

Below are some general Aplo Scope imaging starting conditions as a guide to help you find the right settings for your experiment:  

To configure your workspace in Aplo Scope Control for easy use of the Discovery Kit:

1. Change the filter for the 561nm laser channel to use the “Orange – 595nm” filter, which is optimized for the Cy3B dye. There is no need to modify the filter for the 638nm laser channel; the “Red – 673nm” filter is already optimized for AZ647. Instructions on changing illumination channels can be found here.
2. Optionally, save this workspace for quick access in the future. Instructions on how to save and load workspaces can be found here.

Here are some general considerations when modifying the dSTORM imaging settings for both Nanoimager and Aplo Scope.  

Exposure time
A 30 ms exposure 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 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 dSTORM Cy3B- or AZDye™647-labelled targets. 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, the laser power recommended in the tables above is sufficient to produce robust blinking for both Cy3B- or AZDye™647-labelled targets. However, if using a very stringent illumination angle 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 the laser power, 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 a 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 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. If using a Nanoimager, 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. Once the upload has finished, go to CODI and navigate to the dataset. 
2. If using Aplo Scope, datasets will be automatically updated to CODI. 

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

For data collected on Aplo Scope, it is often advisable to increase the minimum photon count threshold to 1000, and the maximum localisation precision to 15 nm. 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 applies to a majority of targets. 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, or staining temperature. 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 here. 

Pre-extraction is an optional step that can be used to remove a proportion of the soluble cytosolic molecules within a cell before 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 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). 

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.

Whether Pre-Extraction Solution is 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.

The Discovery Kit Blocking Solution efficiently prevents most non-specific protein deposition within the sample. This ensures a low background when using protein-based probes. In cases where nucleic acid probes are used, such as DNA hybridization 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 when 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 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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