Fixation on Histology

Part II, The Evolution of Multiplex Immunofluorescent Methods in Histotechnology


This is Part II of a two-part series on the evolution of multiplex methods.  Read Part I.

Using immunofluorescence has several advantages over brightfield IHC assays.  Immunofluorescence assays facilitate the visualization of multiple marker colocalization within the same cellular compartment.  Confirmation of overlapping signals is confirmed by turning individual channels off or on during visualization of the images.  In contrast to brightfield chromogenic IHC assays, IF assays have a greater dynamic range and allow us to measure changes in signal intensity.  This provides value in studies where the relative distribution of a protein signal is similar across groups and where there may be a decrease in mean intensity post dose with the investigative drug.  

When considering IF approaches, there are several methods available.  Each of them has their benefits and limitations.  First, there are directly conjugated primary antibodies each with a unique fluorescent tag.  This is convenient because the antibodies can be cocktail together and applied in one incubation step.  A significant drawback of this technique is the lack of amplification which makes it difficult, to not feasible, to visualize using traditional fluorescent microscopy.  A second, more viable option, is using anti-species secondary antibodies against the host species of the primary antibody, that are labeled with fluorescent dyes.  This method is useful for high abundance proteins, but we are limited by the number of animal species available as primary antibodies.  Indeed, the most used animal species for primary antibody production are mice and rabbits so it can be difficult to move beyond a dual assay.  An additional benefit of this method is realized in the ability to increase the sensitivity of this approach by using biotinylated secondary antibodies followed by streptavidin conjugated fluorescent dyes.

 A third multiplex IF technique uses secondary antibodies conjugated with HRP enzymes and tyramide signal amplification resulting in deposition of a fluorescent tag at the site of primary antibody binding.  Multiplexing using this technique involves running a single plex assay followed by a stripping procedure followed by the application of additional single plex assays with the stripping technique in between each round of single plex staining.  Although this technique produces a strong specific signal it has a few disadvantages.  Since the staining method requires sequential application of the individual primary antibodies, the time required to run the assay exponentially increases with each additional marker.  Moreover, the order in which the antibodies are applied can have an impact on the success of each marker.  For example, pan cytokeratin is a highly expressed, robust epitope in positive tissues that is not negatively impacted by the repeated stripping method.  For this reason, pan cytokeratin can be applied in the last position of the staining protocol without losing the intensity of expression.  For all antibodies used in this tyramide approach, the successful detection of each target must be empirically tested by placing the target antibody in multiple positions within the assay.  Unfortunately this increases the time required and the costs associated with the assay validation of tyramide assays.  This technique is also limited to five or six antibody targets within a multiplex assay.  This is due to the limited number of fluorescent dyes that are available for use in this assay.  An additional drawback that is often encountered, is the requirement to perform spectral imaging.  The use of spectral unmixing increases the time and complexity of the assay development. 

Despite these challenges, we continued to drive the development of this approach within the clinical histotechnology lab as the benefits outweighed the limitations.  Two alternative approaches to tyramide include technologies that use a that use an antibody/DNA barcoding approach and the second that use primary antibodies directly labeled with proprietary haptens.  The DNA barcoding method, in brief, involves traditional deparaffinization and antigen retrieval followed by incubation with a cocktail containing four primary antibodies each tagged with a unique DNA sequence (barcode).  This is followed by an amplification step that increases the ratio of barcodes per antibody allowing for more complementary probe strands to bind.  The final detection step uses fluorescently tagged (FITC, TRITC, Cy5, Cy7) DNA probes that hybridize with the DNA targets.  At the completion of the assay, a temporary coverslip is applied to the slide then loaded onto the whole slide scanner for image capture.  The protocol may stop here for a four plex assay or be used in a second round of staining and image with an additional application of four new target antibodies resulting in an eight plex assay. 

The second round of the assay requires the use of a mild decoupling solution following the first round and prior to incubating with the new panel of uniquely barcoded antibodies.  Finally, the section is imaged again and the image from the first round of the assay is fused with the image from the second round.  The temporary coverslip is removed, and the section is stained using a routine hematoxylin and eosin stain.  The section is scanned for a third time and the image is used to produce a brightfield image for morphometric evaluation by the pathologist and to provide landmarks for use image analysis.  This produces a “true reference H&E” since it was produced using the same section as the IF assay.  This approach has multiple advantages over the other previously described multiplexing techniques.  Since the antibodies are cocktail together the position of the antibody in the assay is not a cause for concern.  There is no secondary anti-species antibody used in this approach thus avoiding nonspecific background staining due to secondary antibody binding.  The completion of the four plex assay is more than twice as fast as the tyramide 4 plex assay and the image capture does not require spectral unmixing.  In the second multiplex approach using haptens, primary antibodies are directly labeled with a unique hapten.  Up to 6 haptens are available to use in the assay.  Following traditional dewaxing, antigen retrieval and blocks steps, the primary antibodies are applied a cocktail and incubated for 1 hour followed by detection step.  The detection step uses a cocktail of secondary anti-hapten antibodies that are tagged with specific fluorescent dyes that are aligned to the excitation and emission filters that are installed on the users imaging platform.  Like the DNA barcoding approach, this method avoids issues with species anti-species limitations. 

All of the above approaches are compatible with automated staining platforms allowing efficient and reproducible results with minimal tech hands-on time.  Each lab has its unique requirements and infrastructure so each will need to determine which method will meet their needs.  In addition to these approaches, there is an increasing number of specialized staining and/or imaging platforms that are specifically designed to go beyond the 6-8 plex panels using these techniques.  These immerging hybrid platforms allow researchers to go up to approximately 100 markers. Taken together, these multiplex IF approaches will allow us to conserve limited patient biopsy samples and to dive deeper into the complex world of micro-tissue environment. 

Written by:  David Krull, HT(ASCP), QIHC (ASCP)