Getting Started

General Guidelines for Sample Submission to RHTIC

For paraffin embedding:

  • Please label cassettes in pencil
  • Place tissue into cassettes and close cassettes securely. Please make sure the tissue is small enough to move within the cassette (no thicker than a nickel). If it is too large you might need to dissect it into smaller pieces for it to fix properly. Hard objects such as sutures, hair, or lens will affect sectioning and if not the object of the study, need to be removed prior to fixation/ tissue submission to the core. If tissue is very small, please use biopsy cassettes with denser mesh and add sponges to hold tissue in place.
  • Tissue needs to be fixed immediately. Place cassettes into a container large enough to allow the cassettes to move around and fill it with 10% neutral buffered formalin (10%NBF) or 4%PFA. Please use at least 15-20 volumes of formalin for every volume of tissue.
  • Fix tissues at room temperature for 24-48hr.
  • After fixation, remove formalin and replace it with 70% ethanol.
  • Bring your samples to the Histology core in 70% ethanol in a SEALED container (The core no longer accepts samples in inappropriate containers). If you don’t have the right container, please ask us, and we will try to help you find one.

 

For Preparing Frozen Samples:

  • Select a proper size cryomold and label it with permanent marker. Choose a mold larger than your samples. Having enough OCT around the tissue allows for better sample attachment to the chuck and makes sectioning easier. If your sample is sticking out of the mold, the Core staff won’t be able to section it.
  • If your sample is in PBS or some other liquid, try to blot away as much of the liquid as possible before placing tissue in OCT. Equilibrate your sample to OCT at room temperature for 30 seconds in a small container (i.e. 50ml tube cap) before moving it to the molds (lungs need to be perfused with a 1:1 mixture of OCT and PBS to remove air before freezing to ensure good section quality and correct tissue morphology).
  • Add a few drops of OCT to the mold, transfer your sample and orient it correctly in the mold. The side of the sample facing the bottom of the cryomold is the side that will get sectioned first. Tissue should be placed in the center of the mold. If embedding several pieces of tissue in the same block, place them as close to the center as possible.
  • If needed, carefully add more OCT to completely cover the specimen. Avoid bubbles.
  • Using long forceps, place the mold in vapors of liquid nitrogen/dry ice-isopentane slurry and allow OCT to freeze completely. It should take 30 sec-1 min for OCT to harden and become white. It is preferred that fresh samples are frozen in liquid nitrogen vapors.
  • Some samples might require fixation before freezing in OCT. Samples that are pre-fixed should be incubated in 15% sucrose in PBS, and then in 30% sucrose in PBS until they sink in 30% sucrose. Sucrose helps to displace water and to prevent ice crystals formation inside the sample, which results in a better tissue morphology. Pre-fixed samples can be frozen on dry ice.
  • Wrap each mold in aluminum foil to prevent samples from drying and write the sample ID on the foil using permanent marker. Place the block in a labeled zip lock bag. Store samples at -80C and transport them on dry ice.

 

For Slide Scanning:

  • All samples must be clearly labeled, have a coverslip that completely covers the sample, and must be completely dry before submitting to the facility. If using an aqueous-based mounting media, seal the slides with nail polish.
  • Protected Health Information (clinical resection number, patient’s name, ect) is NOT allowed on slide labels and must be covered with a new flat and thin label with a coded sample ID.
  • All slides must be clean. Depending on the type of the mounting media used, use xylene, 70% ethanol, or commercial slides/objectives cleaning solution to remove mounting media residues, fingerprints and dust from the top and the bottom surfaces of the slides.
  • Try your best to avoid bubbles on the sample.  Regions adjacent to bubbles are likely to be out of focus.
  • If preparing fluorescently labeled samples, avoid using mounting media with DAPI. Staining for DAPI separately before putting a coverslip on always produces better results. We recommend ProLong Diamond Antifade Mountant from Molecular Probes.
  • Broken slides can sometimes be manually scanned on the Aperio in brightfield but cannot be scanned on the Vectra.

 

Please use the following link to place an order with RHTIC in iLab:

iLab Link

  • Bring samples to RHTIC together with the printed iLabs-generated order form (please bring histology samples to MSB E311 and samples for scanning and analysis to COMRB 6094).
  • Please allow at least 10 business days (although it can take longer depending on the queue) for your order to be processed. If you need it sooner, please let us know – there will be an additional RUSH charge applied for the expedited service