TITLE: Cell DIVE™ Platform | Antibody Staining & Imaging
AUTHORS: Liz McDonough, Chrystal Chadwick, Fiona Ginty, Christine Surrette, Anup Sood

[DESCRIPTION]
<div class = "text-blocks"><div class = "text-block">This document outlines the workflow for antibody staining, dye inactivation, and imaging as per the Cell DIVE™ technology.</div></div>

[STEPS]
?. [Background Imaging]
Refer to Cell DIVE Imaging Manual.
?. [Decoverslipping]
Coverslips are removed from slides to prepare them for the next slide processing step.
?. [Decoverslipping]
Take a plastic slide box and place it in secondary containment.
?. [Decoverslipping]
Fill the plastic slide box with 1x PBS.
?. [Decoverslipping]
Rest the coverslipped slides on the interior raised edges of slide box such that they are inverted (ie. coverslip down and barcode up).
?. [Decoverslipping]
If your next round is a staining round, prepare your dilutions (see Determining the Order of Antibodies for Staining and Calculations for Antibody Cocktails) while you are waiting for the coverslips to come off.
?. [Decoverslipping]
Be patient - within -, coverslips should naturally come off.
?. [Determining the Order of Antibodies for Staining]
Below is a table of the segmentation markers available for use in the single cell analysis workflow. These markers are required to run single cell analysis.  ABC1Segmentation MarkerRequired for SCA?Segmentation used for2DAPIYNuclear3S6YCytoplasmic4NaKATPaseYMembrane5PCK26YEpithelialThese segmentation markers are critical to the single cell analysis pipeline, and early staining of them is important to avoid the presence of any tissue damage or movement in the images, if the tissue is susceptible to either. That being said, the order of the segmentation markers relies on what other antibodies are in the study. The following rules are recommended for determining the order of antibodies for staining:If there are antibodies being used with primary/secondary detection, those must be stained prior to any direct conjugate raised in the same species. Antibodies that have antigen effects from dye inactivation should be stained in an early round.Once staining with the primary-secondary antibodies and the antibodies with antigen effects is completed, proceed onto the following segmentation markers            o S6            o NaK             o PCK26
ABC1Segmentation MarkerRequired for SCA?Segmentation used for2DAPIYNuclear3S6YCytoplasmic4NaKATPaseYMembrane5PCK26YEpithelial
?. [Calculations for Antibody Cocktails]
The following section should be used when making cocktails of antibodies for any type of experiment including primary/secondary (P/S) antibody, directly conjugated (DC) antibody, or zenon-labelled antibody. For P/S antibodies, the species (Ex- rabbit, mouse IgG1) must be different to run on the same slide at the same time.
?. [Calculations for Antibody Cocktails]
Determine the total volume that will be needed for the experiment.
?. [Calculations for Antibody Cocktails]
Determine the stock concentration for the antibodies. This should be written on the barcode or label on the antibody. If there is no concentration written on the vial or the datasheet, contact either the vendor and or chemist who prepared it and ask. Provide the catalog number and lot#, if applicable.
?. [Calculations for Antibody Cocktails]
Once a concentration is obtained for an antibody, determine the final concentration the antibodies or conjugates should be run at. If the antibody/DC is new and commercially available and no final concentration is noted on the vendor datasheet, call the vendor to ask. If no final concentration can be recommended, run the new marker at .
?. [Calculations for Antibody Cocktails]
Use the following formula to calculate the required amount for each antibody (Ab) -stock concentration*X=final concentration Ab1*total volume needed for experiment stock concentration*X=final concentration Ab2*total volume needed for experiment Solve for X in each case to determine how many µL of each antibody/DC will be required. Subtract the sum of the Ab volumes together from the total volume to get the volume of Ab diluent (3% BSA in PBS) that needs to be added.
For example - if 20 µL of Ab1 and 40 µL of Ab2 was used, the sum of Ab volumes=60 µL. Subtract this volume from the total volume needed for the experiment to obtain the volume of diluent to add to the Abs.
?. [Calculations for Antibody Cocktails]
If you are using secondary antibodies, determine the volume you will need for the secondary in the same manner described above for calculating for primary antibody/DC. Secondary antibodies are stained at a final concentration of . Determine which secondaries you need based on the isotypes of the P/S antibodies used.
?. [Calculations for Antibody Cocktails]
Determine if there is a dye preference for each marker (ie. Some markers look better in Cy3 or Cy5). If there is no preference, place the marker that has lower expression in Cy3 if possible.
Check to make sure the dyes are different when making cocktails with DCs. For P/S antibodies, different species as well as different dyes must be used for each antibody. For example, a mouse IgG1 antibody and a rabbit IgG antibody could be pooled and an example of pooled secondary antibodies to use could be Cy3-Donkey anti-mouse and Cy5- Donkey anti-rabbit.
?. [Calculations for Antibody Cocktails]
If you are staining with -primary/secondary antibodies, click below to proceed to 1°/2° Ab. directly conjugated antibodies, click below to proceed to Conjugated Ab. zenon-labeled antibodies, click below to proceed to Zenon Ab.
?. [Ab Staining – Primary-Secondary Antibodies]
Retrieve primary antibody from the refrigerator or  freezer. Be sure to keep  at all times.
4 °C
-20 °C
on ice
?. [Ab Staining – Primary-Secondary Antibodies]
Spin the stock antibody down at . If stock antibodies are in large or odd shaped vials, transfer them to eppendorf tubes and label appropriately with a LIMS sticker (if possible, remove the LIMS label from the original vial and stick it on the eppendorf tube).
Centrifuge: 13000 34, 1 min
?. [Ab Staining – Primary-Secondary Antibodies]
Make up primary antibody dilutions in the antibody diluent solution () using the manufacturer suggested concentration of antibody, with the exceptions below.If there is no concentration suggested, then use If the vendor suggests , then increase it to  (when performing 1 hour room temperature incubation).
[BSA in 1x PBS]
?. [Ab Staining – Primary-Secondary Antibodies]
Spin down dilutions at  and keep dilutions . Put antibodies back to proper storage location after making up dilutions.
Centrifuge: 13000 34, 1 min
on ice
?. [Ab Staining – Primary-Secondary Antibodies]
Slides should be in 1X PBS (from decoverslipping step); remove from PBS and turn slide on its side and gently tap slide on blue pad/Kimwipe to remove solution. Remove as much of the PBS solution without letting the tissue dry out and without touching the tissue.
?. [Ab Staining – Primary-Secondary Antibodies]
Place slides in humidified chamber. Chamber should contain water in the bottom. Move quickly in adding the primary antibody so that slides do not dry out.
?. [Ab Staining – Primary-Secondary Antibodies]
Add a total of  to each slide; cover with parafilm cut to a size just larger than tissue area. Make sure there are no bubbles under the parafilm. Place lid on humidified chamber.
[antibody dilution]
?. [Ab Staining – Primary-Secondary Antibodies]
Incubate for  at .
0 Room temperature
?. [Ab Staining – Primary-Secondary Antibodies]
Retrieve secondary antibody aliquot from ; keep  and then spin down at .
-20 °C
on ice
Centrifuge: 13000 34, 1 min
?. [Ab Staining – Primary-Secondary Antibodies]
Make up secondary antibody dilutions in the antibody diluent solution (). Secondary antibodies are stained at a final concentration of .
[BSA in 1x PBS]
?. [Ab Staining – Primary-Secondary Antibodies]
Spin down dilutions at . Keep  in light protected environment while the primary antibody incubation is taking place.
Centrifuge: 13000 34, 1 min
on ice
?. [Ab Staining – Primary-Secondary Antibodies]
Remove parafilm and place slides in coplin jar or staining dish. Wash slides 3x at 5 minutes each with 1X PBS, using gentle agitation on shaker:
?. [Ab Staining – Primary-Secondary Antibodies]
(Wash 1/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Ab Staining – Primary-Secondary Antibodies]
(Wash 2/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Ab Staining – Primary-Secondary Antibodies]
(Wash 3/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Ab Staining – Primary-Secondary Antibodies]
After final PBS wash, turn slide on its side and gently tap slide on blue pad/Kimwipe to remove remaining PBS. Once again remove as much of the liquid without letting the tissue dry out and without touching the tissue.
?. [Ab Staining – Primary-Secondary Antibodies]
Place slides in humidified chamber. Add a total of  to each slide just as suggested above. Cover with parafilm cut to a size just larger than tissue area. Make sure there are no bubbles under the parafilm. Place lid on humidified chamber.
[secondary antibody dilution]
?. [Ab Staining – Primary-Secondary Antibodies]
Incubate for  at .
0 Room temperature
?. [Ab Staining – Primary-Secondary Antibodies]
Remove parafilm and place slides in coplin jar or staining dish. Wash slides 3x at 5 minutes each with 1X PBS, using gentle agitation on shaker:
?. [Ab Staining – Primary-Secondary Antibodies]
(Wash 1/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Ab Staining – Primary-Secondary Antibodies]
(Wash 2/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Ab Staining – Primary-Secondary Antibodies]
(Wash 3/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
A 2 minute DAPI recharge is recommended but not required after staining.
?. [Ab Staining – Primary-Secondary Antibodies]
Proceed immediately to Coverslipping.
?. [Coverslipping]
Slides are coverslipped to protect the tissue from drying out.
?. [Coverslipping]
Mounting media should be removed from the fridge and brought to .
4 °C
0 Room temperature
?. [Coverslipping]
Slides should be quickly dipped in ddH2O.
?. [Coverslipping]
Take a rainin pipet tip (200uL-green box) and with a scissors clip the end to make a wide bore.
?. [Coverslipping]
Take up  (4% propyl gallate, 1% DABCO, 90% glycerol) and add to one end of the slide. For leaching markers, use alternate mounting media (50% glycerol, 4% propyl gallate).
[mounting media]
?. [Coverslipping]
Place the end of the coverslip at a 45 degree angle and slowly lower; allowing the mounting media to flow under the coverslip across the slide.
?. [Coverslipping]
Make sure there is not mounting media oozing out of the sides of the coverslip; remove any excess by gently blotting on a Kimwipe.
?. [Coverslipping]
Store the slides at  in light protected environment. When ready for imaging, proceed to Imaging. Recommendation: image within 24 hours of staining.
4 °C
?. [Imaging]
Proceed to Imaging as per the Cell DIVETM Imaging manual.If you are imaging a stain, you are conducting a stain round. If you are imaging post-dye inactivation, you are conducting a background round.
?. [Imaging]
After imaging, you will proceed on to either dye inactivation or staining.If you just imaged your background round and are ready to stain, proceed to Decoverslipping and then  Determining the Order of Antibodies for Staining. If you just imaged your stain ready and are ready to dye inactivate, proceed to Section Decoverslipping and then  Dye Inactivation.
?. [Dye Inactivation]
Dye inactivation is performed after every staining round, except for the last round. Dye inactivation rounds vary for each stain based on the bleaching efficiency, but the majority of rounds will need just 1x of dye inactivation post-stain. (PCK26 and AE1 require 2-3 rounds of dye inactivation.)
?. [Dye Inactivation]
The acceptable range for the pH of the 0.5 M NaHCO3 is between   and  . Check and log the pH on a weekly basis during periods of use. If the pH is measured outside of this specification, discard the buffer and do not use. Refer to the antibody catalogue to understand which antibodies have antigen effects with this dye inactivation method. Use this information to design your multiplexing experiment such that the antigens sensitive to dye inactivation are stained first.
?. [Dye Inactivation]
Fill a “PBS” labeled staining dish with fresh 1X PBS. Place decoverslipped slides to a slide rack and transfer to this staining dish.
?. [Dye Inactivation]
Transfer Dye Inactivation Solution into a “Dye Inactivation” labeled staining dish.
Refer to the working solution for dye inactivation. The hydrogen peroxide is added to the solution immediately before adding to the staining dish. This reaction is time sensitive.
?. [Dye Inactivation]
Transfer the slide rack into “Dye Inactivation” staining dish and incubate for  at .
0 Room temperature
?. [Dye Inactivation]
If more than one round is needed, following each dye inactivation incubation/round, transfer slide rack into staining dish containing fresh 1X PBS for . Prepare a fresh batch of Dye Inactivation Solution for the next round during this one minute wash.
?. [Dye Inactivation]
Once the dye inactivation rounds are complete, wash the slides 3X with fresh 1X PBS for 5 minutes each with gentle agitation on an orbital shaker set to 60 RPM:
?. [Dye Inactivation]
(Wash 1/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
(Wash 2/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
(Wash 3/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
DAPI signal diminishes after dye inactivation. Slides are re-stained with DAPI to label the cell nuclei for imaging purposes.
?. [Dye Inactivation]
Prepare the DAPI Staining Solution.
?. [Dye Inactivation]
Pour the DAPI Staining Solution into a “DAPI” labeled staining dish.
?. [Dye Inactivation]
Transfer the slide rack sitting in 1X PBS into the “DAPI” staining dish.
?. [Dye Inactivation]
Incubate slides in DAPI Staining Solution for  at  with gentle agitation on an orbital shaker set to 60 RPM.
0 Room temperature
?. [Dye Inactivation]
After the 2 minute incubation, transfer slide rack into a staining dish containing fresh 1X PBS.
DO NOT DISCARD DAPI; the DAPI may be reused up to 10 times or by the expiration date, whichever comes first. Tally the number of uses on the DAPI bottle
?. [Dye Inactivation]
Wash slides for  in 1X PBS with gentle agitation on an orbital shaker set to 60 RPM.
?. [Dye Inactivation]
Proceed immediately to Coverslipping and then Imaging.
?. [Ab Staining – Directly Conjugated Antibodies]
Retrieve the directly conjugated Abs from the  walk-in refrigerator or  freezer. Be sure to keep  at all times.
4 °C
-20 °C
on ice
?. [Coverslipping]
Slides are coverslipped to protect the FFPEs from drying out.
?. [Coverslipping]
Mounting media should be removed from the 4C fridge and brought to .
0 Room temperature
?. [Coverslipping]
Slides should be quickly dipped in ddH2O.
?. [Coverslipping]
Take a rainin pipet tip (200uL-green box) and with a scissors clip the end to make a wide bore.
?. [Coverslipping]
Take up  (4% propyl gallate, 1% DABCO, 90% glycerol) and add to one end of the slide. For leaching markers, use alternate mounting media (50% glycerol, 4% propyl gallate).
[mounting media]
?. [Coverslipping]
Place the end of the coverslip at a 45 degree angle and slowly lower; allowing the mounting media to flow under the coverslip across the slide.
?. [Coverslipping]
Make sure there is not mounting media oozing out of the sides of the coverslip; remove any excess by gently blotting on a Kimwipe.
?. [Coverslipping]
Store the slides at  in light protected environment. When ready for imaging, proceed to Imaging. Recommendation: image within 24 hours of staining.
4 °C
?. [Imaging]
Proceed to Imaging as per the Cell DIVE Imaging manual.If you are imaging a stain, you are conducting a stain round. If you are imaging post-dye inactivation, you are conducting a background round.
?. [Imaging]
After imaging, you will proceed on to either dye inactivation or staining.If you just imaged your background round and are ready to stain, proceed to Decoverslipping and then  Determining the Order of Antibodies for Staining. If you just imaged your stain round and are ready to dye inactivate, proceed to Decoverslipping and then Dye Inactivation.
?. [Dye Inactivation]
Dye inactivation is performed after every staining round, except for the last round. Dye inactivation rounds vary for each stain based on the bleaching efficiency, but the majority of rounds will need just 1x of dye inactivation post-stain. (PCK26 and AE1 require 2-3 rounds of dye inactivation.)
?. [Dye Inactivation]
The acceptable range for the pH of the 0.5 M NaHCO3 is between   and  . Check and log the pH on a weekly basis during periods of use. If the pH is measured outside of this specification, discard the buffer and do not use. Refer to the antibody catalogue to understand which antibodies have antigen effects with this dye inactivation method. Use this information to design your multiplexing experiment such that the antigens sensitive to dye inactivation are stained first.
?. [Dye Inactivation]
Fill a “PBS” labeled staining dish with fresh 1X PBS. Place decoverslipped slides to a slide rack and transfer to that dish.
?. [Dye Inactivation]
Transfer Dye Inactivation Solution into a “Dye Inactivation” labeled staining dish.
Refer to the working solution for dye inactivation. The hydrogen peroxide is added to the solution immediately before adding to the staining dish. This reaction is time sensitive.
?. [Dye Inactivation]
Transfer the slide rack into “Dye Inactivation” staining dish and incubate for  at .
0 Room temperature
?. [Dye Inactivation]
Following each dye inactivation incubation/round, transfer slide rack into staining dish containing fresh 1X PBS for .
?. [Dye Inactivation]
Prepare a fresh batch of Dye Inactivation Solution for the next round during this one minute wash. After the solution is prepared, repeat steps 44 and 45 and proceed onto 47:
?. [Dye Inactivation]
Transfer the slide rack into “Dye Inactivation” staining dish and incubate for  at .
0 Room temperature
?. [Dye Inactivation]
If more than one round is needed, following each dye inactivation incubation/round, transfer slide rack into staining dish containing fresh 1X PBS for . Prepare a fresh batch of Dye Inactivation Solution for the next round during this one minute wash.
?. [Dye Inactivation]
Once the dye inactivation rounds are complete, wash the slides 3X with fresh 1X PBS for 5 minutes each with gentle agitation on an orbital shaker set to 60 RPM:
?. [Dye Inactivation]
(Wash 1/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
(Wash 2/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
(Wash 3/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
DAPI signal diminishes after dye inactivation. Slides are re-stained with DAPI to label the cell nuclei for imaging purposes.
?. [Dye Inactivation]
Prepare the DAPI Staining Solution.
?. Pour the DAPI Staining Solution into a “DAPI” labeled staining dish.
?. Transfer the slide rack sitting in 1X PBS into the “DAPI” staining dish.
?. Incubate slides in DAPI Staining Solution for  at  with gentle agitation on an orbital shaker set to 60 RPM.
0 Room temperature
?. After the 2 minute incubation, transfer blue slide rack into a staining dish containing fresh 1X PBS.
DO NOT DISCARD DAPI; the DAPI may be reused up to 10 times or by the expiration date, whichever comes first. Tally the number of uses on the DAPI bottle
?. Wash slides for 5 minutes in 1X PBS with gentle agitation on an orbital shaker set to 60 RPM:
?. (Wash 1/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. (Wash 2/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. (Wash 3/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. Proceed immediately to Coverslipping and then Imaging.
?. [Calculating Ab Amount for Ab Dilution Cocktails]
?. [Calculating Ab Amount for Ab Dilution Cocktails]
?. [Antibody Staining – Zenon Labelled Antibodies]
Retrieve primary antibody from the  walk-in refrigerator or  freezer. Be sure to keep  at all times. Note that the IgG concentration of Ab should be known. If unknown, contact vendor. If no concentration can be supplied, assume 200µg/mL.
4 °C
-20 °C
on ice
?. [Coverslipping]
Slides are coverslipped to protect the tissue from drying out.
?. [Coverslipping]
Mounting media should be removed from the 4C fridge and brought to .
0 Room temperature
?. [Coverslipping]
Slides should be quickly dipped in ddH2O.
?. [Coverslipping]
Take a rainin pipet tip (200uL-green box) and with a scissors clip the end to make a wide bore.
?. [Coverslipping]
Take up  (4% propyl gallate, 1% DABCO, 90% glycerol) and add to one end of the slide. For leaching markers, use alternate mounting media (50% glycerol, 4% propyl gallate).
[mounting media]
?. [Coverslipping]
Place the end of the coverslip at a 45 degree angle and slowly lower; allowing the mounting media to flow under the coverslip across the slide.
?. [Coverslipping]
Make sure there is not mounting media oozing out of the sides of the coverslip; remove any excess by gently blotting on a Kimwipe.
?. [Coverslipping]
Store the slides at  in light protected environment. When ready for imaging, proceed to Imaging. Recommendation: image within 24 hours of staining.
4 °C
?. [Imaging]
Proceed to Imaging as per the Cell DIVE Imaging manual.If you are imaging a stain, you are conducting a stain round. If you are imaging post-dye inactivation, you are conducting a background round.
?. [Imaging]
After imaging, you will proceed on to either dye inactivation or staining.If you just imaged your background and are ready to stain, proceed to Decoverslipping and then Determining the Order of Antibodies for Staining. If you just imaged your stain round and are ready for dye inactivation, proceed to Decoverslipping and then Dye Inactivation.
?. [Dye Inactivation]
Dye inactivation is performed after every staining round, except for the last round. Dye inactivation rounds vary for each stain based on the bleaching efficiency, but the majority of rounds will need just 1x of dye inactivation post-stain. (PCK26 and AE1 require 2-3 rounds of dye inactivation.)
?. [Dye Inactivation]
The acceptable range for the pH of the 0.5 M NaHCO3 is between   and  . Check and log the pH on a weekly basis during periods of use. If the pH is measured outside of this specification, discard the buffer and do not use. Refer to the antibody catalogue to understand which antibodies have antigen effects with this dye inactivation method. Use this information to design your multiplexing experiment such that the antigens sensitive to dye inactivation are stained first.
?. [Dye Inactivation]
Fill a “PBS” labeled staining dish with fresh 1X PBS. Place decoverslipped slides to a slide rack and transfer to that dish.
?. [Dye Inactivation]
Transfer Dye Inactivation Solution into a “Dye Inactivation” labeled staining dish.
Refer to the working solution for dye inactivation. The hydrogen peroxide is added to the solution immediately before adding to the staining dish. This reaction is time sensitive.
?. [Dye Inactivation]
Transfer the slide rack into “Dye Inactivation” staining dish and incubate for  at .
0 Room temperature
?. [Dye Inactivation]
If more than one round is needed, following each dye inactivation incubation/round, transfer slide rack into staining dish containing fresh 1X PBS for . Prepare a fresh batch of Dye Inactivation Solution for the next round during this one minute wash.
?. [Dye Inactivation]
Once the dye inactivation rounds are complete, wash the slides 3X with fresh 1X PBS for 5 minutes each with gentle agitation on an orbital shaker set to 60 RPM:
?. [Dye Inactivation]
(Wash 1/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
(Wash 2/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
(Wash 3/3) Wash slides for  with 1X PBS, using gentle agitation on shaker set to 60 RPM.
?. [Dye Inactivation]
DAPI signal diminishes after dye inactivation. Slides are re-stained with DAPI to label the cell nuclei for imaging purposes.
?. [Dye Inactivation]
Prepare the DAPI Staining Solution.
?. [Dye Inactivation]
Pour the DAPI Staining Solution into a “DAPI” labeled staining dish.
?. [Dye Inactivation]
Transfer the slide rack sitting in 1X PBS into the “DAPI” staining dish.
?. [Dye Inactivation]
Incubate slides in DAPI Staining Solution for  at  with gentle agitation on an orbital shaker set to 60 RPM.
0 Room temperature
?. [Dye Inactivation]
After the 2 minute incubation, transfer blue slide rack into a staining dish containing fresh 1X PBS.
DO NOT DISCARD DAPI; the DAPI may be reused up to 10 times or by the expiration date, whichever comes first. Tally the number of uses on the DAPI bottle
?. [Dye Inactivation]
Wash slides for  in 1X PBS with gentle agitation on an orbital shaker set to 60 RPM.
?. [Dye Inactivation]
Proceed immediately to Coverslipping and then Imaging.
?. [Ab Staining – Directly Conjugated Antibodies]
Spin the stock antibody down at .
Centrifuge: 13000 34, 1 min
?. [Ab Staining – Directly Conjugated Antibodies]
Make up direct conjugate antibody dilutions (as per their validated concentrations) in the antibody diluent buffer () using amber tubes for direct conjugates.
[BSA in 1x PBS]
?. [Ab Staining – Directly Conjugated Antibodies]
Spin the dilutions at  and keep dilutions . Put antibodies and direct conjugates back to proper storage location after making up dilutions.
Centrifuge: 13000 33, 1 min
on ice
?. [Ab Staining – Directly Conjugated Antibodies]
Slides should be in 1X PBS (from decoverslipping step); remove from PBS and turn slide on its side and gently tap slide on blue pad/Kimwipe to remove solution. Remove as much of the PBS solution without letting the tissue dry out and without touching the tissue.
?. [Ab Staining – Directly Conjugated Antibodies]
Place slides in humidified chamber. Chamber should contain water in the bottom. Move quickly in adding the primary antibody so that slides do not dry out.
?. [Ab Staining – Directly Conjugated Antibodies]
Add a total of  to each slide; cover with parafilm cut to a size just larger than tissue area. Make sure there are no bubbles under the parafilm. Place lid on humidified chamber.
[antibody dilution]
?. [Ab Staining – Directly Conjugated Antibodies]
Incubate for  at .
0 Room temperature
?. [Ab Staining – Directly Conjugated Antibodies]
Remove parafilm and place slides in coplin jar or staining dish. Wash slides 3x at 5 minutes each with 1X PBS, using gentle agitation on shaker.
?. [Ab Staining – Directly Conjugated Antibodies]
(Wash 1/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Ab Staining – Directly Conjugated Antibodies]
Proceed immediately to Coverslipping.
?. [Ab Staining – Directly Conjugated Antibodies]
(Wash 2/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Ab Staining – Directly Conjugated Antibodies]
(Wash 3/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
A 2 minute DAPI recharge is recommended but not required after staining.
?. [Antibody Staining – Zenon Labelled Antibodies]
Mix the zenon and antibody target at the validated molar ratio and concentration. Use a pipet to mix (up and down) instead of vortexing due to the low volumes).
?. [Antibody Staining – Zenon Labelled Antibodies]
Incubate the tubes for  at .
0 Room temperature
?. [Antibody Staining – Zenon Labelled Antibodies]
Add an amount of zenon blocking reagent to each tube equivalent to the amount of labeling reagent used (i.e. if you used 10 µL of zenon label, add 10 µL of the zenon blocking reagent).
?. [Antibody Staining – Zenon Labelled Antibodies]
Incubate the tubes for  at .
0 Room temperature
?. [Antibody Staining – Zenon Labelled Antibodies]
Bring the volume of each tube to the desired volume (200 µL per slide * the number of slides) using antibody diluent () for the manual staining.
[BSA in 1x PBS]
Conjugates must be applied to slides within 30 minutes.
?. [Antibody Staining – Zenon Labelled Antibodies]
Slides should be in 1X PBS (from decoverslipping step); remove from PBS and turn slide on its side and gently tap slide on blue pad/Kimwipe to remove solution. Remove as much of the PBS solution without letting the tissue dry out and without touching the tissue.
?. [Antibody Staining – Zenon Labelled Antibodies]
Place slides in humidified chamber. Chamber should contain water in the bottom. Move quickly in adding the primary antibody so that slides do not dry out.
?. [Antibody Staining – Zenon Labelled Antibodies]
Add a total of  to each slide; cover with parafilm cut to a size just larger than tissue area. Make sure there are no bubbles under the parafilm. Place lid on humidified chamber.
[antibody dilution]
?. [Antibody Staining – Zenon Labelled Antibodies]
Incubate for  at .
0 Room temperature
?. [Antibody Staining – Zenon Labelled Antibodies]
Remove parafilm and place slides in coplin jar or staining dish. Wash slides 3x at 5 minutes each with 1X PBS, using gentle agitation on shaker:
?. [Antibody Staining – Zenon Labelled Antibodies]
Proceed immediately to Coverslipping.
?. [Antibody Staining – Zenon Labelled Antibodies]
(Wash 1/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Antibody Staining – Zenon Labelled Antibodies]
(Wash 2/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
?. [Antibody Staining – Zenon Labelled Antibodies]
(Wash 3/3) Wash slides for  with 1X PBS, using gentle agitation on shaker.
A 2 minute DAPI recharge is recommended but not required after staining.
?. [Bleaching]
Pour the DAPI Staining Solution into a “DAPI” labeled staining dish.
?. [Bleaching]
Transfer the slide rack sitting in 1X PBS into the “DAPI” staining dish.
?. [Bleaching]
Incubate slides in DAPI Staining Solution for  at  with gentle agitation on an orbital shaker set to 60 RPM.
0 Room temperature
?. [Bleaching]
After the 2 minute incubation, transfer blue slide rack into a staining dish containing fresh 1X PBS.
DO NOT DISCARD DAPI; the DAPI may be reused up to 10 times or by the expiration date, whichever comes first. Tally the number of uses on the DAPI bottle
?. [Bleaching]
Wash slides for  in 1X PBS with gentle agitation on an orbital shaker set to 60 RPM.
?. [Bleaching]
Proceed immediately to the coverslipping, Section "Coverslipping", and then imaging, Section "Imaging".
?. [Analysis]
Please refer to the following manuals for analysis –Understanding the images from acquisition: Section 7 in the Cell DIVE Imaging Manual Viewing the imagines in the Visualization Software: Layers Visualization Manual Analyzing the images with Single Cell Analysis: Image Analysis Manual Analyzing the results from the Single Cell Analysis: Data Analysis Manual
?. [Antibody Staining – Zenon Labelled Antibodies]