HOPE tissue fixation

Kindly provided by DCS - Innovative Diagnostik- Systeme

For your own safety:

The HOPE technique is used for the preparation of paraffin embedded tissue sections. In contrast to other fixation methods HOPE does not completely denature structural proteins, enzymes, and nucleic acids. They remain in a nearly native state. This means that HOPE-fixed tissue can also include aktive virus, prions, microorganisms etc. HOPE-fixed tissue blocks, paraffin sections etc. therefore must be considered as potentially infectous unless other suitable tests have proven the opposite. Always wear gloves!

HOPE reagents contain up to 0,03 % NaN3 (sodium azide). Sodium azide is not classified as an hazardous chemical at the concentration of these products. However, toxicity information about sodium azide at the products' concentration has not been thoroughly investigated. For further information, a Material Safety Data Sheet for sodium azide in pure form is available upon request.

1.) Tissue Fixation and Preparation of Paraffin Blocks

  • Take tissue in OR and transfer directly into sterile plastic petri dish and cool on wet ice. Take care that the tissue does not dry out. Seal the dish with Parafilm if necessary. DO NOT transfer into physiological salt solution. As soon as HOPE I solution is available, cut tissue into pieces of max. 8 mm x 8 mm x 3 mm and transfer into disposable 5 ml tubes (must withstand acetone!) with ice-cold (0-4°C) HOPE I solution.
  • Important: To ensure good diffusion and penetration of the tissue, only use cut/sectioned organs and organ parts, respectively, i.e. do NOT use encapsulated tissues! If using frozen tissue, do NOT thaw prior to transfer into HOPE I solution. Be aware of the possibility of lesser-quality morphology, if working with frozen tissue.
  • Store tissue in HOPE I solution at 0-4°C for 1 - 3 nights (i.e. 16 - 64 hours). During this incubation, erythrocytes might diffuse out of the tissue and form a reddish pellet in the tube. However, this does not affect the quality of the fixation. The color of HOPE I solution should stay orange-yellow throughout the fixation representing a stable pH.
    For processing of tissue samples with high cell density (like brain or lymph nodes) incubation in HOPE I solution for at least 2 nights (about 40 hours) should be preferred.
    NOTE: Samples are still potentially infectious - always wear gloves!
  • In the morning, empty tube carefully (tissue has to stay inside the tube!). If necessary, place tube upside down on tissue to empty completely. Add 5 ml pre-mixed ice-cold acetone solution (100 ml acetone + 100 µl HOPE II solution) to tissue in tube and transfer immediately back into fridge or ice bath. Ignore possibly visible white precipitate. Incubate at 0-2°C for 2 hours.
    NOTE: The quality of the fixation and the performance during cutting of the paraffin-embedded HOPE-fixed tissue is critically dependent on the acetone not being warmer than 2°C. To ensure this, please, perform the dehydration with acetone in an ice bath and ensure it remains around freezing temperature throughout the incubation time of 8 hours total.
  • After 2 hours, empty tube again (compare above) and re-fill with ice-cold pure acetone. Incubate for 2 hours. Repeat twice (each incubation: 2 hours).
  • After 8 hours of dehydration (Do not dehydrate longer than for 8 hours total), empty acetone out of tube (Careful: Tissue is still potentially infectious!) and immediately add 5 ml pre-warmed low-melting paraffin. Make sure tissue does not dry out in between steps. Try to get rid of all air bubbles. Incubate overnight at exactly 54-55°C.
  • Paraffinizing of tissue is complete the next morning. Incomplete penetration of tissue at this point cannot be helped by longer incubation. Reembed tissue as usual using fresh paraffin. Avoid any air bubbles. Basically proceed as with formalin-fixed tissue. Chill blocks immediately on ice.
    NOTE: Cracks in the paraffin blocks occur more often than with regular paraffin due to the absolutely pure paraffin used here.
    Dispose used paraffin and 5 ml tubes.
  • Store blocks in fridge until sectioning.
  • Remarks for transportation of tissue samples for HOPE fixation: Technical Protocols for transport of tissue material for HOPE fixation are still under development. The best way should be to transfer the tissue immediately into cold HOPE I solution and incubate for at least 16 hours. If, during the subsequent transport, the tissue stored in HOPE I becomes slightly warmer, no negative impact on morphology was observed up to now.

2.) Sectioning of HOPE-Fixed Paraffin-Embedded Tissue

Please, give yourself time to get a little practice cutting HOPE-fixed tissue training with less important material!

  • Store paraffin blocks at about -20°C for 30 min and fix on microtome as usual. After all other usual preparations are finished, the temperature of the blocks will have risen so that sectioning is possible without cracking the tissue. Prepare two different water baths and set #1 at RT, #2 at 35 - 39°C. Before you start sectioning prepare two cuvettes with pure isopropanol and keep in a 60°C incubator.
  • Transfer sections onto the surface of water bath #1 (RT) without dipping it into the water. You can collect your sections here. Fish sections with clean slide and, one by one, stretch them out on the surface of water bath #2 (35 - 39°C). Make sure sections do not dissolve. As soon as section is stretched out and wrinkle-free on slide, take out of water bath, drip-dry carefully on tissue and place on drying rack. Dry sections in dry incubator at 50°C for about 30 minutes or (preferably) at 37°C overnight.
  • Dried sections can be stored in the fridge without problems for a long peroid of time. Blocks should be stored at 4 °C.
  • For deparaffinization, place slides into first cuvette with 60°C isopropanol. Incubate for 10 minutes. Transfer to second cuvette with 60°C isopropanol and wash thoroughly. Drip-dry slide on tissue and air-dry. Slides are now deparaffinized and can be stored in the fridge, if desired, but be aware of the risk of the tissue picking up moisture there.
  • Rehydrate tissue by incubating it in 70% ice-cold acetone for a minimum of 10 minutes in the fridge. Take slides out of the acetone, drip-dry on tissue (5 seconds) and transfer the still moist slide into a cuvette with aqua dest. Wash thoroughly. Transfer to a second batch of aqua dest. and incubate for 10 minutes. Briefly drip-dry slides and transfer onto a hot plate set to 45°C until complete evaporation of the water. Tissue should be dried onto the slide at this point (takes about 1-2 minutes on the hot plate).
  • To stain with H & E, transfer re-hydrated section to hematoxylin for 2-4 minutes. Wash rigorously in a cuvette with aqua dest and transfer to a second cuvette with aqua dest. Perform bluing reaction under running tap water (1-2 minutes). Important: Avoid any influence of acids!
    Incubate in eosin for 2-4 minutes depending on the intensity desired. Wash twice in aqua dest and dehydrate rapidly dipping the slide into the following solutions:
    2x 70% Isopropanol
    2x Isopropanol abs. Incubate for another 10 minutes in a third cuvette with absolute Isopropanol. Briefly wash in xylene or Rotihistol, then transfer into a second batch of xylene or Rotihistol and incubate for 5 minutes.
  • Take out, drip-dry and coverslip.
  • Notes re. IHC: In general, blocking with serum is not necessary. NEVER block endogenous peroxidase in MetOH solution with H2O2, since this is highly likely to destroy antigen structures (epitopes). Instead, simply block using buffer solutions or water with 0.3 - 0.5% peroxide. Dissolve antibodies and enzyme conjugates in PBS or TBS only. Use same dilution as in frozen tissue.
  • Notes re. ISH: If at all possible, avoid SDS and dextran sulfate in hybridization mix. Enzymatic digestion is hardly ever necessary.

3.) Immunohistochemistry (IHC) on HOPE-Fixed Tissue Sections

This chapter describes two methods for IHC on HOPE-fixed tissue sections. In general all IHC systems suitable for formalin-fixed and frozen sections will also work on HOPE-fixed material, but there are several advantages and drawbacks from one system to the other.


  • Place deparaffinized and rehydrated tissue sections (preparation in chapter 2) in a cuvette with 0.5 % H2O2 in DPBS (= PBS according to Dulbecco without Ca2+ and Mg2+) for 30 minutes to block endogenous peroxidase. Re.: peroxidase-based detection systems are recommended, since endogenous alkaline phosphatase activity in Hope-fixed tissues will very often not be quenched completely even when using levamisole. After H2O2 block wash slides in a cuvette with aqua dest and transfer immediately into a new cuvette with DPBS.
  • Take slides out of DPBS, dip dry briefly and remove excess liquid on the backside of the slide as well as around the section with tissue. (Do not allow sections to dry out during the whole staining procedure!) Apply primary antibody diluted in DPBS without serum, albumins etc. Dilution of the primary antibody is about the same as for frozen sections. In general, protein block with sera is not necessary but should be tested by each lab.
    Below we describe an IHC protocol using a mouse primary antibody.
  • After 60 minutes incubation with the primary antibody at room temperature (longer incubation times never improved the results on HOPE-fixed tissue) rinse slides with DPBS, wash briefly in a cuvette with DPBS and incubate in a second cuvette with DPBS for about 2 minutes.
  • Take slides out of DPBS, remove excess liquid (see above) and cover sections with a digoxigenin-conjugated anti-mouse secondary antibody diluted in DPBS and incubate for 60 minutes at room temperature. (Re: This protocol uses an digoxigenin/anti-digoxigenin system. If you want to work with biotin/streptavidin on HOPE-fixed material, we recommend to test a biotin block). Incubate secondary antibody for 60 minutes at room temperature.
  • Wash slides as described above and apply a tertiary antibody (e.g. anti-digoxigenin peroxidase-conjugate) diluted in DPBS. Incubate for 60 minutes at room temperature.
  • Wash slides and apply substrate/chromogen (e.g. AEC or DAB) for approx. 10 -20 minutes. Rinse slides with aqua dest, counterstain and coverslip with synthetic aqueous mounting medium (e.g. Aquatex, Merck # 1.08562). Counterstain of nuclei seems to be less stable in glycerol gelatine, therefore synthetic mounting media should be preferred.

PROTOCOL 2 (short protocol):

This method is based on the streptavidin/biotin technique and corresponds to the standard detection principle in routine IHC on formalin-fixed tissue sections. The method has been tested with excellent results, although the number of different tissue types tested up to now is small.

As mentioned in protocol 1, use of streptavidin/biotin systems on HOPE-fixed tissue can sometimes result in a higher background staining. If the level of background is inacceptable suitable blocking steps or biotin-free detection systems should be preferred.

  • To block endogenous peroxidase, incubate deparaffinized and rehydrated tissue sections (preparation see chapter 2) for 10 minutes in a cuvette with 0.3 % H2O2 (in aqua dest). Wash slides with PBS. (Note: Do not allow sections to dry out during the whole staining procedure!).
  • Take slides out of PBS and perform a serum block for 10 minutes to reduce background staining. Drain slides, do not wash.
  • Apply primary antibody for 60 minutes at room temperature. The primary antibody should be diluted in PBS without serum, albumin etc.. Dilution should be approx. 3-fold compared to formalin-fixed sections.
  • Wash slides in PBS and incubate with biotinylated secondary antibody (diluted in PBS, e.g. DCS PolyLink secondary antibody, anti-mouse, anti-rabbit, anti-rat) for 30 minutes at room temperature. Dilution of the secondary should be approx. 3-fold compared to formalin-fixed tissue sections.
  • Wash slides in PBS and apply peroxidase label (diluted in PBS, e.g. DCS streptavidin peroxidase conjugate) for 30 min at room temperature. Dilution of the peroxidase label should be approx. 3-fold compared to formalin-fixed tissue sections.
  • Wash slides in PBS and cover with substrate/chromogen (e.g. DCS AEC-1-step-solution or DAB-1-step-solution). Incubate for appx. 10 minutes at room temperature. Wash slides with aqua dest, counterstain and coverslip with synthetic aqueous mounting medium (e.g. Aquatex, Merck # 1.08562). Counterstain of nuclei seems to be less stable in glycerol gelatine, therefore synthetic mounting media should be preferred.


Please keep in mind that HOPE-fixed tissue samples are potentially infectious! HOPE-fixed tissue can include active virus, microorganisms etc. For your own safety always wear gloves!

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