Multiplexed antibody-based imaging enables the characterization of cellular interactions, spatial relationships, and histological patterns in tissues. These techniques require considerable resources and domain expertise for implementation, limiting their widespread adoption. To overcome these challenges, we developed Iterative Bleaching Extends multi-pleXity (IBEX), an open-source method that can be completed at relatively low cost in 2-5 days by biologists with basic laboratory skills. In addition to cost-effective techniques such as IBEX, community initiatives are needed to extend the reach of multiplexed tissue imaging. The IBEX Imaging Community is an international group of scientists committed to sharing knowledge related to multiplexed imaging in a transparent and collaborative manner. Organ Mapping Antibody Panels (OMAPs) are community-validated resources that save time and money, increase reproducibility, and support the construction of a Human Reference Atlas. This webinar will provide guidelines for the successful application of IBEX to a wide range of scientific questions using diverse imaging platforms. A significant amount of time will be devoted to the following topics: optimal tissue processing, working with fixed frozen and FFPE specimens, identifying appropriate reagents, and constructing organ atlases. Team science initiatives will be highlighted to encourage attendees to join and accelerate discovery together.

Multiplexed antibody-based imaging empowers the study of complex cellular phenotypes in situ by enabling the evaluation of dozens of protein biomarkers in a single tissue section. Iterative Bleaching Extends multi-pleXity (IBEX), is an antibody-based staining and chemical bleaching technique that allows comprehensive profiling (65+ parameters) in diverse tissues. It is compatible with 250+ commercially available antibodies and 16 unique fluorophores. It can be adopted by different imaging platforms at a low cost, and the experiment can be done in 2-5 days by biologists with basic laboratory skills. In this webinar, we will go over methods to study normal and malignant lymph nodes and topics related to multiplexed tissue imaging, namely: optimal tissue processing, working with fixed frozen and FFPE specimens, overcoming autofluorescence, identifying appropriate reagents, validating antibodies, and creating organ mapping antibody panels (OMAPs).

After the slide (containing coverslip) has been soaked in 1X PBS pH 7.4 to allow coverslip to loosen (typically 30-60 minutes), carefully remove slide from container. Ideally coverslip will have fallen off naturally. If coverslip is loose but still attached, gently lift a corner of the coverslip with a pair of fine forceps or razor blade.

We typically wait until coverslip detaches from slide naturally or, at minimum, is moving freely on the slide under the force of gravity. Attempting to remove coverslip before it has sufficiently loosened may result in tissue deformation and/or loss.

Dissolve LiBH₄ (STREM Chemicals, cat no. 93-0397; purchase in 1 g aliquots) into diH₂O (Quality Biological, cat no. 351-029-101) and allow to sit at RT for 10 minutes until the formation of bubbles occurs (not shown in video). Pass the solution through a 0.22 μm syringe filter (Millipore Sigma, cat no. SLGSM33SS) attached to a 20 mL syringe (EXEL Int., cat no. 26280) to remove any impurities prior to use (not shown).

Always prepare LiBH₄ solution in a chemical fume hood with appropriate personal protective equipment (PPE). Mixing LiBH₄ with diH₂O will generate hydrogen gas which is highly flammable. For this reason, we always work with small amounts of LiBH₄ relative to the amount of water initially added. We do not mix >10 mg of LiBH₄ with diH₂O. This amount can be quickly diluted to 1 mg/mL with volumes handled by standard serological pipets/pipet-aids. Always wrap LiBH₄ stock with parafilm (suggest Bemis Company Inc., cat no. S37440) and store in the presence of desiccant. We recommend replacing LiBH₄ every 4 weeks as repeated exposure to moisture/air has been observed to reduce fluorophore inactivation efficacy. Be sure to follow institutional guidelines for proper disposal of hazardous chemicals.

The formation of bubbles indicates the solution is ready for filtration and subsequent use. For best results, use solution immediately after filtration. We have noted effective fluorophore inactivation up to 4 hours after initial dissolution of LiBH₄.

Coat a glass slide by adding 15 μL chrome alum-gelatin to one side. Spread evenly using the edge of a cover slip or a separate glass slide via the blood smear technique. Note, it may require multiple spreading passages with the slide to ensure even coating of chrome alum-gelatin without streaks. If chrome alum-gelatin pools on the slide, aspirate excess fluid and respread as these accumulations can result in autofluorescence artifacts during image acquisition. Be mindful of stated expiration date on chrome alum-gelatin.

For best results, prepare chrome alum-gelatin coated slides freshly (i.e., the day of tissue sectioning) or no more than 7 days prior to tissue sectioning. If chrome alum-gelatin coated slides are prepared ahead of sectioning, store at room temperature (do not freeze as this will compromise the adhesive properties).

Dry coated slide in oven at 60 degrees Celsius for 1 hour.

We purchase chrome alum-gelatin from Newcomer Supply, but recipes are available online if this item does not ship to your country (e.g. this recipe).

Remove as much PBS as possible without drying out tissues. Quickly add the minimum amount of Fluoromount-G mounting medium necessary to completely cover each tissue section. Gently cover with a coverslip. We typically use 10-40 μL per tissue section. Ensure there are no bubbles on or near tissue.

This video illustrates how to use the XTRegisterSameChannel SimpleITK Imaris (Oxford Instruments) Python Extension - registration of 2D or 3D images that share a common channel (correlation based affine alignment).

This is a video tutorial on how to set up a focus map in LAS X Navigator (v. 3.6.0 Widefield), which is recommended for tissue sections or samples that are continuous.

How to use Adaptive Focus Control (AFC) in your Navigator experiments, if your DMi8 microscope is equipped with AFC.

A video tutorial on how to do Linked Shading (shading correction) in LAS X 3.6 (widefield) with fluorescent images on a monochrome camera.

This video shows how to "THUNDER" images in LAS X on-the-fly or post-acquisition with the default settings. (v. 3.7.1 - Widefield). This is only available on THUNDER Imagers.

How to set up a Z-stack in LAS X v 3.3 (widefield). Note: the example in this video was for a data set that was intended for deconvolution, so it goes slightly beyond the focus on either end. The usual recommendation is to stay mostly in focus for all of your Z planes, so that subsequent 2D projections (such as a maximum intensity projection or extended depth of focus projection) will be clearer.