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Nov 2015

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Fluorescent Labeling of Rat-tail Collagen for 3D Fluorescence Imaging
用于3D荧光成像的鼠尾胶的荧光标记   

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Abstract

Rat tail collagen solutions have been used as polymerizable in vitro three-dimensional (3D) extracellular matrix (ECM) gels for single and collective cell migration assays as well as spheroid formation. These 3D hydrogels are a relatively inexpensive, simple to use model system that can mimic the in vivo physical characteristics of numerous tissues within the body, namely the skin. While confocal imaging techniques such as fluorescence reflection and two-photon microscopy are able to visualize collagen fibrils during 3D imaging without fluorescence, other imaging modalities require direct conjugation of fluorescent dyes to collagen. Here we detail how to generate 3D collagen gels labeled with a fluorescent dye. Furthermore, we go through the steps required to reproducibly generate bright collagen hydrogels that are suitable for live cell 3D imaging techniques.

Keywords: Rat-tail collagen (鼠尾胶), Fibrils (原纤维), Hydrogel (水凝胶), Fluorescent dye (荧光染料), 3D imaging (3D成像)

Background

The study of cell migration and cell interaction with its surrounding microenvironment has been started since the 1950’s when Paul Weiss and Beatrice Garber originally observed the effect of increasing plasma concentration (fibrin) on mesenchymal cell morphology (Weiss and Garber, 1952). In subsequent years and decades, biochemists started to delve into purifying extracts from rat tail collagen and started their use as a highly polymerizable 3D matrix (Fitch et al., 1955; Gross et al., 1955; Chandrakasan et al., 1976). It wasn’t until the 1990’s that 3D matrices truly became useful to the cell biology community, especially for studying cell migration (Friedl et al., 1995). Recently, a transition from simplified two-dimensional (2D) studies on ECMs to 3D has begun. This evolution has followed shortly behind the recent advances in fluorescence microscopy, especially super-resolution microscopy. While standard laser scanning confocal microscopes can utilize reflection microscopy or two-photon-based second harmonic generation to visualize collagen fibers in the absence of a fluorescent tag, both techniques do not always properly depict the ECM architecture due either to polarity issues or lack of a significant fibril thickness. A fluorescently-tagged ECM allows imaging of the smallest individual fibrils even with super-resolution techniques.

Unlike most proteins collagen cannot be simply tagged with a fluorophore when in solution because the numerous lysine residues are required for alpha helix formation with other monomers during polymerization (Chandrakasan et al., 1976). For this reason, the labeling must be accomplished on preformed gels. This protocol describes how to label a polymerized gel, bring the collagen back into solution with acetic acid, and properly mix a minimal amount (2-4% of total protein) of labeled collagen with an unlabeled fraction to generate a bright, fluorescent collagen gel capable of sustaining cell viability while allowing observation of ECM architecture over multiple hours of fluorescence imaging.

Materials and Reagents

  1. MatTek Dishes (35 mm, #1.5 coverslip, 20 mm opening: MATTEK, catalog number: P35G-1.5-20-C )
  2. ColorpHast pH-indicator-strips with pH range 6.5-10 (Merck, catalog number: 109543 )
  3. 10-100 and 1,000 μl Gilson MICROMAN® positive displacement pipette tips (Gilson, catalog numbers: FD10004 , FD10006 )
  4. 10 cm tissue culture dish (Thermo Fisher Scientific, catalog number: 150350 )
  5. Cell lifters (Corning, catalog number: 3008 )
  6. Aluminum foil
  7. Plastic wrap
  8. Scintillation vial (Sigma-Aldrich, catalog number: Z190527 )
  9. 1.5 ml microfuge tubes (Thermo Fisher Scientific, catalog number: AM12400 )
  10. Dubecco’s Minimal Essential Medium (DMEM) powder, Phenol red (Sigma-Aldrich, catalog number: D2429 )
  11. NaOH pellets (Sigma-Aldrich, catalog number: S8045 )
  12. Phosphate Buffered Saline with Calcium and Magnesium (PBS++) chilled to 4 °C and at room temperature (GE Healthcare, HycloneTM, catalog number: SH30264.02 )
  13. Rat tail collagen solution (dissolved in 20 mM acetic acid, commercial brands are fine, but in-house preparations are usually cleaner and polymerize faster) at a concentration greater than 5 mg/ml (6 mg/ml used here)
  14. Sodium bicarbonate (Sigma-Aldrich, catalog number: S5761 )
  15. 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (Sigma-Aldrich, catalog number: H4034 )
  16. Boric acid (powder 99.5%, Sigma-Aldrich, catalog number: B6768 )
  17. Acetic acid (Sigma-Aldrich, catalog number: 27221-1L ) (chilled to 4 °C, less than 50 ml of 16.65 M is required)
  18. Atto-488 NHS-ester 1 mg (Sigma-Aldrich, catalog number: 41698 )
  19. DMSO (Sigma-Aldrich, catalog number: D2650 )
  20. Sircol Collagen Assay kit (available from Accurate Chemical & Scientific, catalog number: CLRS1000 )
  21. Slide-A-Lyzer Dialysis cassette (G2) with 20,000 MW cutoff (Thermo Fisher Scientific, catalog number: 87735 )
  22. 1 M Tris (pH 7.4, KD Medical, catalog number: RGF-3340 )
  23. NaCl (Sigma-Aldrich, catalog number: S7653 )
  24. HCl (37%, Sigma-Aldrich, catalog number: H1758-500ML )
  25. 10x DMEM (see Recipes)
  26. 10x reconstitution buffer (10x RB; see Recipes)
  27. 1 N NaOH (500 μl in a microfuge tube; see Recipes)
  28. 1 N HCl (500 μl in a microfuge tube; see Recipes)
  29. 50 mM Borate buffer pH 9.0 (see Recipes)
  30. 5 mg/ml Atto-488 NHS-ester dye in DMSO (see Recipes)
  31. Sodium Chloride (NaCl), 8% solution (see Recipes)
  32. 50 mM Tris buffer (see Recipes)

Equipment

  1. 10-100 and 1,000 μl Gilson MICROMAN® positive displacement pipette (Gilson, catalog numbers: F148314 , F148180 )
  2. Lab timer
  3. 2-4 L beaker
  4. Ultra-clear 8 x 20 mm centrifuge tubes (Optional: Beckman Coulter, catalog number: 345843 )
  5. Rectangular ice bucket packed with ice
  6. Water bath
  7. Large Magnetic stir bar to large beaker (Sigma-Aldrich, SP Scienceware - Bel-Art Products - H-B Instrument, catalog number: Z284491 )
  8. Small Magnetic stir bar for scintillation vial (Sigma-Aldrich, SP Scienceware - Bel-Art Products - H-B Instrument, catalog number: Z126942 )
  9. Stir plate
  10. Mini centrifuge (i.e., VWR, model: Galaxy Mini )
  11. Biological hood (any brand)
  12. Brightfield microscope with 10 and 20x phase contrast objectives
  13. Cooled centrifuge capable of 20,000 x g and 4 °C (i.e., TOMY, model: MX-307 )
  14. Lab Rocker (i.e., Denville Scientific, model: 110, catalog number: S2110 )
  15. Airfuge ultra centrifuge (optional: Beckman Coulter, catalog number: 340400 )
  16. Laser scanning or spinning disk confocal microscope (i.e., Nikon Instruments, model: A1R or Yokogawa, model: CSU-X1 )

Procedure

Notes: If you are planning on repeatedly making collagen gels, it is highly recommended to invest in a 10-100 and a 100-1,000 μl positive displacement pipette. These greatly help accurate pipetting of viscous solutions such as collagen. The greater your accuracy in pipetting, the better the overall experimental reproducibility.

  1. Collagen gel calculations
    1. Prior to any collagen gel generation, it is important to accurately calculate the volumes of the different components needed from stock solutions. Because only a small fraction (between 2-4%) of labeled collagen is used for gel generation, a relatively small amount of collagen is used for the labeling process. Typically, a heavy week of experiments will entail generating 20-24 dishes containing 150 μl of collagen and use approximately 2,000 μl of stock concentration collagen. This amounts to a range of 40 to 80 μl or 120 to 240 μg of labeled collagen being used per week. Here, the 5 ml of labeled 3 mg/ml collagen should last between 37 and 75 weeks of similar usage. Smaller volumes of collagen can be labeled, but it is recommended to use no less than 2.5 ml due to normal losses in the protocol. The following steps walk you through the initial calculations.
      1. Calculate the amount of collagen:



      2. Divide by 10 to calculate the amount of 10x RB and 10x DMEM :

        10x DMEM:
        10x RB:

      3. The volume of 1 N NaOH will vary based on the pH of your stock collagen concentration which can vary between 1.8 and up 3.0. A good starting point is to add 3 μl per ml of collagen, then adjust accordingly. We find 16 μl is suitable for our homemade collagen.
        Note: This will vary greatly depending on the original pH of your collagen and can even vary between commercial lots and preparations. Initial testing may be required for determining the actual amount needed per dish.
      4. Calculate the amount of PBS++:



  2. Collagen Gel preparation
    1. Pre-chill all gel components on ice (collagen stock solution, 10x DMEM, 10x RB, 1 N NaOH, 1 N HCl, PBS++) and the 10 cm tissue culture dish. Add the tissue culture dish last: be sure no water condenses on the inner dish surface prior to adding gel components.
      Note: To reduce collagen solution loss when transferring from a conical tube to the 10 cm tissue culture dish we find it helpful to mix directly in the dish.
    2. Pipette the proper calculated amount of stock collagen solution (2.5 ml) into the pre-cooled tissue culture dish. Be sure to keep the dish on ice.
    3. Add the 10x DMEM (0.25 ml) and then the 10x RB (0.25 ml). After adding each slowly triturate with the 1 ml positive displacement pipette, making sure not to introduce any bubbles. Alternatively, you can use a cell scraper to homogenize.
    4. Add the calculated 1 N NaOH to the solution and triturate/mix with a positive displacement pipette or cell scraper. Once the solution is mixed well and shows a consistent, single color take a 2 μl sample and test the pH with a pH strip, waiting 1-2 min to determine the pH. The pH should be between 7.0 and 7.4, and the solution color should be peach (pinkish orange: Figure 1). If the solution is over a pH of 7.4, adjust with 1 N HCl, 1 μl at a time. Be sure to note the changes and adjust the PBS++ accordingly including for the 2 μl sample (i.e., if 2 μl of HCl were added, reduce the PBS++ by the same amount).
    5. Add PBS++ (adjusted if necessary) to the tissue culture dish until fully mixed and place again in the ice.
    6. Allow gravity to help spread the collagen evenly over the dish by tilting the dish to 45-degree and rotating the dish.
    7. Cover and place the dish on the bench top and allow the collagen to polymerize at room temperature (approximately 21 °C). Check the collagen gel by eye using a 10x or 20x phase contrast objective on an inverted brightfield microscope after 30 min and then every fifteen minutes thereafter. Small, intertwined fibrils should be visible (Figure 1). Again, the polymerization process can vary preparation to preparation based on the collagen used. Too short an incubation time can lead to loss of collagen gel integrity and an overall loss of collagen monomers and will result in a greatly reduced end-product concentration. Longer incubations have no major effect (always side on longer incubations). Testing in small 150 μl batches is suggested if you are unsure. Also, taking a brightfield image before and after adding the PBS++ will help determine if the structure has dramatically changed.


      Figure 1. Fluorescent-labeling of polymerized collagen gels workflow. Schematic workflow for Procedure B and C of the protocol. Image shows 3 mg/ml rat tail collagen polymerized at 21 °C for 1 h (taken with a 20x objective). Note the appearance of discernable collagen fibrils.

  3. Collagen gel labeling with NHS-ester dye
    1. Once the solution is completely polymerized and formed a gel, add 10 ml of 50 mM borate buffer (pH 9.0) and incubate for 15 min at room temperature.
    2. Meanwhile, calculate the amount of dye needed to properly label the amount of protein within the gel using the following equation:



    3. The above equation is for 1 mg of Atto-488 NHS-ester diluted in 200 μl (5 mg/ml) of DMSO using a 2-molar excess which is recommended by the company.
      Note: Each dye has a different molar-excess that works the best for NHS-conjugation. Do not assume the above will work for all dyes. Over labeling can lead to issues with gel formation later on.
    4. Add 45.28 μl of Atto-488 NHS-ester dye to a 15 ml conical tube and bring the volume up to 5 ml with 50 mM borate buffer and vortex quickly.
    5. Carefully aspirate the borate buffer from the tissue culture dish (bring the dish to a 45-degree angle and siphon off at the bottom edge with an aspiration pipette).
    6. Add the dye solution to the collagen gel and wrap the culture dish with aluminum foil to protect from light. Allow the dye to conjugate to the collagen gel for 1 h at room temperature or 4 h at 4 °C (can do overnight) while rocking.
      Note: At room temperature the majority of the dye will conjugate within the first 20 min.
    7. Aspirate dye and add 10 ml of 50 mM Tris buffer (pH 7.5) to quench the dye reaction. Incubate with rocking for 10 min. Keep the gel covered with foil.
    8. Add 10 ml of PBS++. Rinse gel with PBS++ 6 x over the next 4 h to wash out the excess dye.

  4. Liquefying the collagen gel into a solution
    1. Aspirate PBS++ and invert dish with one side raised on its lid in a tissue culture hood for 10-15 min to reduce the amount of fluid within the gel.
      Note: The remaining steps in this section should be performed at 4 °C.
    2. Add 500 to 1,000 μl of 500 mM acetic acid to the gel. Bring the gel into a cold room and rock slowly for 1 h.
      Note: The larger the volume of acetic acid you add the easier it is to get the gel to go into solution. However, this will decrease your final concentration. We suggest starting with 500 μl and adding extra incrementally over time if needed.
    3. After 1 h, use a cell scraper to mix the gel. Scrape gel to one side of the dish (on an angle) and pipette up gel solution with a 1,000 μl positive displacement pipette set to 750 μl. If using a regular pipette, you may need to cut off the pipette tip at about the 100 μl mark for a larger tip opening because of the gel viscosity.
    4. Transfer the collagen solution to a scintillation vial wrapped in aluminum foil and add a small magnetic stir bar. Stir gel at 4 °C overnight. Check periodically to see if the gel has gone into solution. You should not see any ‘chunks’ of polymerized collagen. If you do, add more acetic acid followed by further stirring at 4 °C for several hours. Alternatively, take 5 to 10 μl and smear this in a 35 mm tissue culture dish and check for solution consistency under a microscope at 10x magnification.
    5. Check the volume of the gel. Based on the starting volume (5 ml) and concentration (3 mg/ml) you can make an educated guess at the concentration. If you need a more concentrated solution proceed to the salt precipitation protocol (Procedure F). If the guesstimate is fine, then continue the next step below.

  5. Dialyzing the dye-labeled collagen solution
    1. Pre-cool 8 L of 20 mM acetic acid at 4 °C. This requires 9.6 ml of a 16.65 M solution (standard purchased concentration).
    2. Add 4 L of 20 mM acetic acid to a large beaker. Wet the Slide-A-Lyzer G2 cassette in the acid. Pipette the collagen into the cassette being sure to remove any air bubbles before closing.
    3. Add a large stir bar and the cassette containing the labeled collagen to the beaker and cover with plastic wrap and use a rubber band to firmly secure the plastic wrap. Cover the upper ¾ of the beaker with aluminum foil to protect the labeled collagen from excess light. Stir for 4 h.
    4. Change acetic acid once and let dialyze further overnight.
    5. Remove collagen solution from the cassette to several 1.5 ml centrifuge tubes. Place tubes in a cooling centrifuge. Spin at 20,000 x g for 1 h at 4 °C. Remove and save the supernatant, being careful not to pull up any of the pellet.
    6. Follow the Sircol collagen assay to determine the collagen’s concentration.
    7. Fluorescently labeled collagen can be stored at 4 °C protected from light for more than one year or indefinitely at -80 °C.

  6. Concentration of collagen solutions using salt precipitation (modified from Chandrakasan et al., 1976)
    1. Perform all steps at 4 °C.
    2. Estimate the current volume of the collagen to be concentrated (should still be in the scintillation vial) and add an equal volume of 8% NaCl.
      Note: A precipitate should begin forming in several minutes.
    3. Stir the solution at 4 °C for 4 h or overnight.
    4. Once the liquid portion is clear with no apparent collagen remaining, transfer the collagen to a 15 ml centrifuge tube.
    5. Spin down the solution at 13,000 x g for 20 min at 4 °C.
    6. Aspirate the supernatant and keep the pellet. Add 2 ml of 500 mM acetic acid (approximately concentrating by two times). 
    7. Add a small magnetic stir bar to the 15 ml tube and stir the solution at 4 °C until collagen goes into solution, 3-4 h.
    8. Go to Procedure E for dialysis.

  7. Calculating and mixing labeled rat tail collagen with unlabeled collagen and getting the proper amounts
    Note: Many researchers have mixed fluorescently-labeled and unlabeled collagen solutions together at specific ratios (1:5, 1:4, etc.) in order to generate a fluorescently visible collagen gel capable of sustaining cell life. Collagen that is completely labeled will not polymerize and often is too bright for actual imaging. Too often the final collagen concentration is incorrectly determined because ratios do not take into consideration differences in collagen concentration between labeled and unlabeled fractions. Furthermore, batch-to-batch differences in the labeled collagen concentration make gel consistency an issue that can alter your experimental outcome. Here is described how to calculate and mix a 2-4% (4% shown) fluorescently-labeled gel based on protein weight to make up a 6 ml volume. Even if the change is minor, we suggest always using the calculated final concentration shown in step 6 below. Note, always keep collagen solutions at 4 °C or on ice to decrease the possibility of protein degradation.
    1. Calculate what 4% of the unlabeled collagen (ULC) is for the amount you want to mix:



    2. Multiple X by 0.04 to get 4% of this amount (Y).



    3. Use this number to then calculate the volume (V) you need to take out from the unlabeled.



    4. Perform the same for calculation for the fluorescently labeled collagen (FLC) as you did in #3 above.



    5. Mix these together in a 15 ml conical tube slowly over 1 h at 4 °C. Transfer to 1.5 ml centrifuge tubes and spin at 20,000 x g for 30 min to remove any debris.
    6. Calculate the ‘new’ amount (C; in mg) since the volumes removed and added in 3 and 4 will likely not be the same.



      Note: Protein (in mg) should be identical to Step G1.
      Then divide the new amount C by the total volume (TV = [total UL RTC – removed] + [added F RTC]).



    7. Label the tube with the concentration and store at 4 °C.

  8. Collagen clearing (Optional)
    Note: To rid the collagen solution of excess debris, either from the original collagen processing or the labeling process, spinning at high RPMs using a Beckman Coulter Airfuge can rapidly clear the solution.
    1. Using positive displacement pipettes, add 450 μl of 4% labeled collagen to four 8 x 20 mm centrifuge tubes.
    2. Add tubes to the centrifuge rotor, lock the device and initiate the spin. Adjust the pressure to register between 25-30 PSI (approximately 90,000-10,000 rpm). Set the time for 30 min.
    3. After 30 min, a colored streak will often be noticeable at the bottom edge of the centrifuge tubes. Being sure not to disturb the pelleted collagen, remove the supernatant to a 15 ml conical tube and store at 4 °C. 
    4. Repeat the process with the remaining collagen (this usually requires a total of 12 tubes).
    5. This 4% fluorescently-labeled collagen can now be used for generating individual collagen gels for live cell microscopy using the calculations in Procedure I (below) and the protocol used in Procedure B.

  9. Collagen gels for 20 mm diameter MatTek glass-bottomed dish
    Note: For a 20 mm diameter glass-bottomed dish 150 μl of collagen solution will generate a 300 μm-thick gel. Due to the high viscosity of collagen solutions and some loss with a small volume/tube add 20 μl extra for each dish for calculations. Always polymerize a minimum of 1 extra dish over what you need. Below is an example calculation of 2 dishes per condition and at a final concentration of 3 mg/ml.
    1. 2 (dishes) x 170 (μl collagen per dish) = 340 μl of collagen solution
    2. Calculate the amount of collagen:



    3. Divide by 10 to calculate the amount of 10x RB and 10x DMEM:

      10x DMEM:
      10x RB:

    4. The volume of 1 N NaOH: ~1 μl 
    5. Calculate the amount of PBS++:



    6. Follow Procedure B. Changing the polymerization temperature will alter the polymerization time and the architecture (Doyle et al., 2015). Decreasing the polymerization temperature leads to a slower gel formation. We hypothesize that decreasing the temperature leads to fewer nucleation sites from which fibrils polymerize, hence the fibrils extend more and create larger pore sizes observed in Figure 2.
    7. It is expected that following this protocol individual fibrils will be easily observable using laser scanning confocal, spinning disk confocal (Figure 2), or light sheet microscopy.


      Figure 2. Fluorescence images of a 4% labeled collagen. A. Single confocal slice (XY) and a 10 μm YZ projection (right) of a 4% atto 488-labeled 3 mg/ml rat-tail collagen gel polymerized at 21 °C. B. 3D rendering of the collagen gel shown in panel A (X, Y, Z dimensions: 100 x 100 x 80 μm, respectively). C. High-resolution max intensity projections of an HT-1080 fibrosarcoma cancer cell transfected with EGFP-α-actinin (magenta) migrating through Atto 565-labeled 3 mg/ml rat-tail collagen (green) polymerized at 16 °C. Images show first frame (left) and 63rd frame (right) in a time-series (imaged every 0.5 μm in Z over 15 microns, every 30 sec). Time is in minutes.

Notes

While using rat tail collagen is a simple and easy way to use 3D hydrogel, it does require repetition to get gels to become consistent. Collagen type I 3D gels is highly dependent on 1) collagen concentration, 2) the ionic concentration, 3) pH, and of course, 4) temperature. Because each of these can affect the time to polymerization, which inevitably leads to the differences in ECM architecture, it is vital that each of the above parameters is consistent between experiments. pH is by far the most variable since it needs to be tested and adjusted to neutralize each batch of collagen made. It is suggested that pH should be checked for each batch and do not assume that the amount of 1 N NaOH will always be the same. Small aliquots of stocks solutions are used throughout the protocol to cut down on changes in concentration due to evaporation that can occur over time.
Other critical issues to be aware of are with the 10x DMEM solution: Due to the high salinity, a precipitate will always be present at 4 °C or when on ice. Trituration of the 10x DMEM solution is important to keep a consistent ionic concentration.

Recipes

  1. 10x DMEM (50 ml)
    1. Add 1 powdered DMEM with phenol red (Sigma-Aldrich) packet into 50 ml distilled water
    2. Stir mixture with heat (approximately 50 °C) until DMEM powder goes into solution
    3. Sterile filter while warm. Make four 10 ml and twenty 0.5 ml aliquots. The 0.5 ml size aliquots are for daily experiments
    4. Store both sizes at -20 °C until use. Upon defrosting heat aliquots in a 37 °C water bath, then vortex and cool on ice
    5. Can be kept indefinitely at -20 °C and 1 month at 4 °C
  2. 10x reconstitution buffer (100 ml)
    1. 2.2 g Sodium bicarbonate (Sigma-Aldrich)
    2. 4.8 g HEPES (Sigma-Aldrich) -or- 20 ml 1 M HEPES stock solution for 0.2 M final
    3. Distilled water up to 100 ml
    4. Filter sterilize and store at -20 °C in aliquots similar to 10x DMEM
    5. Can be kept indefinitely at -20 °C and 1 month at 4 °C
  3. Sodium Hydroxide solution (NaOH), 1 N
    1. 0.5 g NaOH pellets (Sigma-Aldrich)
    2. Distilled water to 12.5 ml
    3. Mix well and filter sterilize, divide into 500 μl aliquots and store indefinitely at -20 °C
  4. 1 N HCl
    1 ml HCl (37% or 12 M, Sigma-Aldrich)
    Distilled water to 12 ml
    Filter sterilize, divide into 500 μl aliquots and store indefinitely at -20 °C
  5. 50 mM Borate buffer (pH 9.0)
    1. 1.55 g boric acid (powder 99.5%, Sigma-Aldrich)
    2. Add distilled water to 400 ml
    3. Add several solid NaOH pellets (Sigma-Aldrich) at a time while mixing until the pH is ~9.0
    4. Add distilled water to 500 ml
    5. Filter sterilize using a 0.2 μm filter
    6. Store up to 1 year at room temperature
  6. 5 mg/ml fluorescent NHS-ester dye in DMSO (200 μl)
    1. Atto-488 NHS-ester 1 mg (Sigma-Aldrich)
      Note: Any NHS-ester dye is suitable and can be substituted.
    2. Add 200 μl DMSO (Sigma-Aldrich) to dye tube and mix (wrapped in foil) on a rotational mixer for 1 h
    3. Use immediately and store excess in 10-20 μl aliquots
    4. Store at -80 °C
  7. Sodium Chloride (NaCl), 8% solution
    1. Dissolve 40 g NaCl (Sigma-Aldrich) in 500 ml distilled water, mix well
    2. Filter sterilize using a 0.2 μm filter
  8. 50 mM Tris buffer (pH 7.4)
    5 ml 1 M Tris (KD Medical)
    Add distilled water to 100 ml
    Filter sterilize using a 0.2 μm filter
    Store up to 1 year at room temperature

Acknowledgments

I would like to thank Joshua Collin, Gavrel Pacheco, and Tomoko Ikeuchi for their helpful comments. This work was supported by the NIDCR Division of Intramural Research. This work was adapted from previous work (Doyle, 2016). The author declares no conflicts of interest.

References

  1. Chandrakasan, G., Torchia, D. A. and Piez, K. A. (1976). Preparation of intact monomeric collagen from rat tail tendon and skin and the structure of the nonhelical ends in solution. J Biol Chem 251(19): 6062-6067.
  2. Doyle, A. D., Carvajal, N., Jin, A., Matsumoto, K. and Yamada, K. M. (2015). Local 3D matrix microenvironment regulates cell migration through spatiotemporal dynamics of contractility-dependent adhesions. Nat Commun 6: 8720.
  3. Doyle, A. D. (2016). Generation of 3D collagen gels with controlled diverse architectures. Curr Protoc Cell Biol 72: 10 20 11-10 20 16.
  4. Fitch, S. M., Harkness, M. L. and Harkness, R. D. (1955). Extraction of collagen from tissues. Nature 176(4473): 163.
  5. Friedl, P., Noble, P. B., Walton, P. A., Laird, D. W., Chauvin, P. J., Tabah, R. J., Black, M. and Zanker, K. S. (1995). Migration of coordinated cell clusters in mesenchymal and epithelial cancer explants in vitro. Cancer Res 55(20): 4557-4560.
  6. Gross, J., Highberger, J. H. and Schmitt, F. O. (1955). Extraction of collagen from connective tissue by neutral salt solutions. Proc Natl Acad Sci U S A 41(1): 1-7.
  7. Weiss, P. and Garber, B. (1952). Shape and movement of mesenchyme cells as functions of the physical structure of the medium: contributions to a quantitative morphology. Proc Natl Acad Sci U S A 38(3): 264-280.

简介

大鼠尾胶原溶液已被用作可聚合的体外三维(3D)细胞外基质(ECM)凝胶,用于单一和集体细胞迁移测定以及球状体形成。 这些3D水凝胶是相对便宜,易于使用的模型系统,其可以模拟体内许多组织(即皮肤)的体内物理特征。 虽然诸如荧光反射和双光子显微镜的共焦成像技术能够在没有荧光的3D成像期间可视化胶原原纤维,但是其他成像模式需要荧光染料直接缀合到胶原。 在这里,我们详细介绍了如何生成用荧光染料标记的3D胶原凝胶。 此外,我们还经历了可重复生成适用于活细胞3D成像技术的明亮胶原水凝胶所需的步骤。

【背景】自20世纪50年代以来,Paul Weiss和Beatrice Garber最初观察到增加血浆浓度(纤维蛋白)对间充质细胞形态的影响(Weiss和Garber,1952),开始研究细胞迁移和细胞与周围微环境的相互作用。在随后的几十年中,生物化学家开始深入研究从鼠尾胶原中纯化提取物,并开始将其用作高度可聚合的3D基质(Fitch et al。,1955; Gross et al。,1955; Chandrakasan et al。,1976)。直到20世纪90年代,3D矩阵才真正对细胞生物学界有用,尤其是研究细胞迁移(Friedl et al。,1995)。最近,从简化的ECM二维(2D)研究到3D的转变已经开始。这种演变紧随荧光显微镜最近的进展,特别是超分辨率显微镜。虽然标准激光扫描共聚焦显微镜可以利用反射显微镜或基于双光子的二次谐波生成来在没有荧光标签的情况下显现胶原纤维,但这两种技术并不总是正确地描绘ECM结构,因为极性问题或缺乏显着的原纤维厚度。荧光标记的ECM即使使用超分辨率技术也可以对最小的单个原纤维进行成像。

与大多数蛋白质不同,胶原蛋白在溶液中不能简单地用荧光团标记,因为在聚合过程中与其他单体形成α螺旋需要大量的赖氨酸残基(Chandrakasan et al。,1976)。因此,标签必须在预制凝胶上完成。该协议描述了如何标记聚合凝胶,将胶原蛋白带回醋酸溶液中,并将少量(2-4%总蛋白)标记胶原蛋白与未标记的部分适当混合,生成明亮的荧光胶原凝胶能够维持细胞活力,同时允许在多个小时的荧光成像中观察ECM结构。

关键字:鼠尾胶, 原纤维, 水凝胶, 荧光染料, 3D成像

材料和试剂

  1. MatTek菜肴(35毫米,#1.5盖玻片,20毫米开口:MATTEK,目录号:P35G-1.5-20-C)
  2. ColorpHast pH指示条,pH范围6.5-10(默克,目录号:109543)
  3. 10-100和1,000μlGilsonMICROMAN ®正位移移液器吸头(Gilson,目录号:FD10004,FD10006)
  4. 10cm组织培养皿(Thermo Fisher Scientific,目录号:150350)
  5. 细胞提升器(康宁,目录号:3008)
  6. 铝箔
  7. 保鲜膜
  8. 闪烁瓶(Sigma-Aldrich,目录号:Z190527)
  9. 1.5 ml微量离心管(Thermo Fisher Scientific,目录号:AM12400)
  10. Dubecco的最小必需培养基(DMEM)粉末,酚红(Sigma-Aldrich,目录号:D2429)
  11. NaOH颗粒(Sigma-Aldrich,目录号:S8045)
  12. 磷酸盐和镁的磷酸盐缓冲盐水(PBS ++ )冷却至4°C并在室温下(GE Healthcare,Hyclone TM ,目录号:SH30264.02)< br />
  13. 大鼠尾胶原蛋白溶液(溶于20 mM乙酸,商业品牌很好,但内部制剂通常更清洁,聚合更快)浓度大于5 mg / ml(此处使用6 mg / ml)
  14. 碳酸氢钠(Sigma-Aldrich,目录号:S5761)
  15. 4-(2-羟乙基)-1-哌嗪乙磺酸(HEPES)(Sigma-Aldrich,目录号:H4034)
  16. 硼酸(粉末99.5%,Sigma-Aldrich,目录号:B6768)
  17. 乙酸(西格玛奥德里奇,目录号:27221-1L)(冷藏至4°C,低于50毫升,16.65米是必需的)
  18. Atto-488 NHS-酯1mg(Sigma-Aldrich,目录号:41698)
  19. DMSO(Sigma-Aldrich,目录号:D2650)
  20. Sircol Collagen Assay试剂盒(可从Accurate Chemical&amp; Scientific获得,目录号:CLRS1000)
  21. Slide-A-Lyzer透析盒(G2),截止频率为20,000 MW(Thermo Fisher Scientific,目录号:87735)
  22. 1 M Tris(pH 7.4,KD Medical,目录号:RGF-3340)
  23. NaCl(Sigma-Aldrich,目录号:S7653)
  24. HCl(37%,Sigma-Aldrich,目录号:H1758-500ML)
  25. 10x DMEM(见食谱)
  26. 10x重构缓冲液(10x RB;参见食谱)
  27. 1N NaOH(微量离心管中500μl;参见配方)
  28. 1 N HCl(微量离心管中500μl;参见配方)
  29. 50 mM硼酸盐缓冲液pH 9.0(参见食谱)
  30. 在DMSO中的5mg / ml Atto-488 NHS-酯染料(参见配方)
  31. 氯化钠(NaCl),8%溶液(见食谱)
  32. 50 mM Tris缓冲液(见食谱)

设备

  1. 10-100和1,000μlGilsonMICROMAN ®容积式移液器(Gilson,目录号:F148314,F148180)
  2. 实验室计时器
  3. 2-4 L烧杯
  4. 超透明8 x 20 mm离心管(可选:Beckman Coulter,目录号:345843)
  5. 装满冰的长方形冰桶
  6. 水浴
  7. 大型烧杯大型磁力搅拌棒(Sigma-Aldrich,SP Scienceware - Bel-Art Products - H-B Instrument,目录号:Z284491)
  8. 用于闪烁瓶的小磁力搅拌棒(Sigma-Aldrich,SP Scienceware - Bel-Art Products - H-B Instrument,目录号:Z126942)
  9. 搅拌盘
  10. 小型离心机(即,VWR,型号:Galaxy Mini)
  11. 生物罩(任何品牌)
  12. 具有10倍和20倍相位对比度的明场显微镜
  13. 冷却离心机能够达到20,000 g和4°C(即,TOMY,型号:MX-307)
  14. Lab Rocker(即。,Denville Scientific,型号:110,目录号:S2110)
  15. Airfuge超级离心机(可选:Beckman Coulter,目录号:340400)
  16. 激光扫描或旋转盘共聚焦显微镜(即,尼康仪器,型号:A1R;或横河电机,型号:CSU-X1)

程序

注意:如果您计划反复制作胶原蛋白凝胶,强烈建议您投资10-100和100-1,000μl容积式移液器。这些有助于准确移取粘性溶液(如胶原蛋白)。移液准确度越高,整体实验重现性越好。

  1. 胶原蛋白凝胶计算
    1. 在产生任何胶原凝胶之前,准确计算储备溶液所需的不同组分的体积是很重要的。因为仅有一小部分(2-4%之间)的标记胶原用于凝胶生成,所以相对少量的胶原用于标记过程。通常,重周的实验将需要产生含有150μl胶原的20-24个培养皿并使用约2,000μl的原液浓度的胶原蛋白。这相当于每周使用40至80μl或120至240μg标记胶原的范围。这里,5ml标记的3mg / ml胶原蛋白应该在相似的使用期间持续37至75周。可以标记较小体积的胶原蛋白,但由于方案中的正常损失,建议使用不少于2.5ml的胶原蛋白。以下步骤将引导您完成初始计算。
      1. 计算胶原蛋白量:

      2. 除以10来计算10x RB和10x DMEM的数量:
        10x DMEM:
        10x RB:
      3. 1N NaOH的体积将根据您的原料胶原浓度的pH值而变化,其浓度可在1.8和3.0之间变化。一个好的起点是每毫升胶原蛋白添加3微升,然后相应调整。我们发现16μl适合我们自制的胶原蛋白。
        注意:这取决于胶原蛋白的原始pH值,并且甚至可以在商业批次和制剂之间变化。确定每道菜所需的实际数量可能需要进行初步测试。
      4. 计算PBS ++ 的数量:


  2. 胶原蛋白凝胶制剂
    1. 将所有凝胶组分在冰上预冷(胶原原液,10x DMEM,10x RB,1N NaOH,1N HCl,PBS ++ )和10cm组织培养皿。最后添加组织培养皿:在添加凝胶成分之前,确保没有水凝结在内盘表面上。
      注意:为了减少从锥形管转移到10厘米组织培养皿时胶原蛋白溶液的损失,我们发现直接在培养皿中混合是有帮助的。
    2. 将适当计算量的原料胶原溶液(2.5ml)移液到预冷的组织培养皿中。一定要把菜放在冰上。
    3. 加入10倍DMEM(0.25毫升),然后加入10倍RB(0.25毫升)。用1ml容量移液管加入各缓慢研磨后,确保不引入任何气泡。或者,您可以使用细胞刮刀进行均质化。
    4. 将计算的1N NaOH加入溶液中,用正位移移液管或细胞刮刀研磨/混合。一旦溶液充分混合并显示一致的单一颜色,取2μl样品并用pH条测试pH,等待1-2分钟以确定pH。 pH值应介于7.0和7.4之间,溶液颜色应为桃色(粉红橙色:图1)。如果溶液的pH值超过7.4,则用1N HCl调节,每次1μl。一定要注意变化,并相应地调整PBS ++ ,包括2μl样品(即,如果加入2μlHCl,则减少PBS ++ 相同数量)。
    5. 将PBS ++ (必要时调整)加入组织培养皿中直至完全混合并再次置于冰中。
    6. 通过将培养皿倾斜45度并旋转培养皿,允许重力帮助将胶原蛋白均匀地铺展在培养皿上。
    7. 将培养皿盖住并放置在工作台顶部,让胶原蛋白在室温下(约21°C)聚合。 30分钟后在倒置明场显微镜上使用10x或20x相衬物镜通过眼睛检查胶原凝胶,然后每15分钟检查一次。应该看到小而交织的原纤维(图1)。同样,聚合过程可以根据使用的胶原改变制剂的制备。孵育时间太短会导致胶原蛋白凝胶完整性的丧失和胶原蛋白单体的整体损失,并导致最终产品浓度大大降低。较长时间的孵化没有重大影响(总是在较长的孵化时间)。如果您不确定,建议以150μl的小批量进行测试。此外,在添加PBS ++ 之前和之后拍摄明场图像将有助于确定结构是否发生了显着变化。



      图1.聚合胶原凝胶工作流程的荧光标记。协议程序B和C的原理图工作流程。图像显示3mg / ml大鼠尾胶原在21℃聚合1小时(用20x物镜拍摄)。注意可辨别的胶原纤维的外观。

  3. 用NHS-酯染料标记胶原蛋白凝胶
    1. 溶液完全聚合并形成凝胶后,加入10 ml 50 mM硼酸盐缓冲液(pH 9.0),在室温下孵育15 min。
    2. 同时,使用以下等式计算正确标记凝胶内蛋白质量所需的染料量:

    3. 上述等式是对于1mg的Atto-488 NHS-酯,用200μl(5mg / ml)DMSO稀释,使用2摩尔过量,这是该公司推荐的。
      注意:每种染料都有不同的摩尔过量,最适合NHS结合。不要认为上述内容适用于所有染料。过度贴标可能会导致凝胶形成问题。
    4. 将45.28μlAtto-488 NHS-酯染料加入15 ml锥形管中,用50 mM硼酸盐缓冲液将体积调至5 ml,并快速涡旋。
    5. 小心地从组织培养皿中吸出硼酸盐缓冲液(将培养皿置于45度角,并用吸管吸取底部边缘的虹吸管)。
    6. 将染料溶液加入胶原凝胶中,用铝箔包裹培养皿,避免光照。让染料在室温下与胶原凝胶结合1小时或在4°C下结合4小时(可以过夜),同时摇摆。
      注意:在室温下,大部分染料将在前20分钟内结合。
    7. 吸出染料并加入10ml 50mM Tris缓冲液(pH7.5)以淬灭染料反应。摇动孵育10分钟。保持凝胶覆盖箔。
    8. 加入10毫升PBS ++ 。在接下来的4小时内用PBS ++ 6 x冲洗凝胶以洗掉过量的染料。

  4. 将胶原凝胶液化成溶液
    1. 吸出PBS ++ 并翻转培养皿,将其一侧盖在组织培养罩中盖上10-15分钟,以减少凝胶中的液体量。
      注意:本节中的其余步骤应在4°C下执行。
    2. 向凝胶中加入500至1,000μl的500mM乙酸。将凝胶放入冷室并缓慢摇动1小时。
      注意:添加的醋酸量越大,凝胶越容易进入溶液状态。但是,这会降低您的最终浓度。我们建议从500μl开始,如果需要,随着时间的推移逐渐增加。
    3. 1小时后,使用细胞刮刀混合凝胶。将凝胶刮到培养皿的一侧(以一定角度)并用1,000μl正位移移液管吸取凝胶溶液,设置为750μl。如果使用常规移液器,由于凝胶粘度,您可能需要在大约100μl标记处切断移液器吸头以获得更大的尖端开口。
    4. 将胶原蛋白溶液转移到用铝箔包裹的闪烁瓶中,并加入小磁力搅拌棒。在4℃下搅拌凝胶过夜。定期检查凝胶是否已进入溶液状态。你不应该看到聚合胶原蛋白的任何'块'。如果这样做,添加更多的乙酸,然后在4°C下进一步搅拌几个小时。或者,取5至10μl并在35 mm组织培养皿中涂抹,并在显微镜下以10x放大倍数检查溶液浓度。
    5. 检查凝胶的体积。根据起始体积(5 ml)和浓度(3 mg / ml),您可以对浓度进行有根据的猜测。如果您需要更浓缩的溶液,请继续进行盐沉淀方案(程序F)。如果猜测没问题,那么继续下面的下一步。

  5. 透析染料标记的胶原溶液
    1. 在4℃下预冷8L 20mM乙酸。这需要9.6毫升16.65 M溶液(标准购买浓度)。
    2. 将4L 20mM乙酸加入大烧杯中。在酸中润湿Slide-A-Lyzer G2盒。将胶原蛋白移入盒式磁带,确保在关闭之前清除任何气泡。
    3. 在烧杯中加入一个大的搅拌棒和装有标记胶原蛋白的盒子,盖上保鲜膜,用橡皮筋牢牢固定保鲜膜。用铝箔覆盖烧杯的上部3/4,以保护标记的胶原蛋白免受过量光照。搅拌4小时。
    4. 改变醋酸一次,再过夜透析。
    5. 将胶原蛋白溶液从盒中取出至几个1.5ml离心管中。将管置于冷却离心机中。在4℃下以20,000 x g 旋转1小时。取出并保存上清液,小心不要拔出任何颗粒。
    6. 按照Sircol胶原蛋白测定法测定胶原蛋白的浓度。
    7. 荧光标记的胶原蛋白可以在4°C下避光保存一年以上或无限期保存在-80°C。

  6. 使用盐沉淀浓缩胶原蛋白溶液(改编自 Chandrakasan 等,1976)
    1. 在4°C下执行所有步骤。
    2. 估计要浓缩的胶原蛋白的当前体积(应该仍然在闪烁瓶中)并添加等体积的8%NaCl。
      注意:沉淀物应在几分钟内开始形成。
    3. 将溶液在4°C搅拌4小时或过夜。
    4. 一旦液体部分澄清,没有明显的胶原蛋白残留,将胶原蛋白转移到15ml离心管中。
    5. 在13,000 x g 下将溶液在4℃下旋转20分钟。
    6. 吸出上清液并保持沉淀。加入2毫升500毫升乙酸(大约浓缩两次)。&nbsp;
    7. 在15 ml管中加入一个小磁力搅拌棒,在4°C下搅拌溶液,直至胶原蛋白进入溶液,3-4小时。
    8. 转到程序E进行透析。

  7. 用未标记的胶原蛋白计算和混合标记的鼠尾胶原蛋白并获得适当的量
    注意:许多研究人员将荧光标记和未标记的胶原蛋白溶液以特定比例(1:5,1:4等)混合在一起,以产生能够维持细胞寿命的荧光可见胶原凝胶。完全标记的胶原蛋白不会聚合,并且通常对于实际成像来说太亮。通常最终的胶原蛋白浓度不正确地确定,因为比率没有考虑标记和未标记部分之间胶原浓度的差异。此外,标记胶原蛋白浓度的批次间差异使凝胶一致性成为可能改变实验结果的问题。这里描述了如何计算和混合基于蛋白质重量的2-4%(显示4%)荧光标记的凝胶以构成6ml体积。即使变化很小,我们建议始终使用下面步骤6中显示的计算的最终浓度。注意,始终将胶原蛋白溶液保持在4°C或冰上,以减少蛋白质降解的可能性。
    1. 计算4%的未标记胶原蛋白(ULC)是否适合您想要混合的量:

    2. 多个 X 按0.04获得此金额的4%( Y )。

    3. 使用此数字然后计算您需要从未标记的卷中取出的音量( V )。

    4. 对上述#3中的荧光标记胶原蛋白(FLC)进行计算也是如此。

    5. 将它们一起在15ml锥形管中在4℃下缓慢混合1小时。转移至1.5 ml离心管并以20,000 x g 旋转30分钟以去除任何碎片。
    6. 计算'新'量( C ;以mg为单位),因为在3和4中删除并添加的卷可能不相同。

      注意:蛋白质(mg)应与步骤G1相同。
      然后将新的 C 除以总体积(TV = [总UL RTC - 删除] + [添加F RTC])。

    7. 用浓度标记管,并在4°C下储存。

  8. 胶原蛋白清除(可选)
    注意:要从原始胶原蛋白处理或标记过程中去除多余碎片的胶原蛋白溶液,使用Beckman Coulter Airfuge以高RPM旋转可快速清除溶液。
    1. 使用正位移移液管,将450μl4%标记的胶原蛋白添加到4个8 x 20 mm离心管中。
    2. 将管加入离心机转子,锁定设备并启动旋转。调节压力以在25-30 PSI(约90,000-10,000 rpm)之间进行调节。设定时间为30分钟。
    3. 30分钟后,在离心管的底部边缘经常会出现彩色条纹。确保不要打扰沉淀的胶原蛋白,将上清液移至15毫升锥形管中,并在4℃下储存。&nbsp;
    4. 用剩余的胶原重复该过程(这通常需要总共12个管)。
    5. 这种4%荧光标记的胶原蛋白现在可用于生成用于活细胞显微术的单个胶原凝胶,使用程序I(下文)中的计算和程序B中使用的方案。

  9. 胶原蛋白凝胶为20毫米直径的MatTek玻璃底盘
    注意:对于20 mm直径的玻璃底皿,150μl胶原蛋白溶液将产生300μm厚的凝胶。由于胶原蛋白溶液的高粘度和一些体积/管的损失,每个培养皿额外增加20μl用于计算。始终聚合至少1个额外的菜超过你需要的。以下是每种条件下2个培养皿的实例计算,最终浓度为3 mg / ml。
    1. 2(餐具)x 170(每个培养皿中的μl胶原蛋白)=340μl的胶原蛋白溶液
    2. 计算胶原蛋白量:

    3. 除以10来计算10x RB和10x DMEM的数量:
      10x DMEM:
      10x RB:
    4. 1N NaOH的体积:~1μl&nbsp;
    5. 计算PBS ++ 的数量:

    6. 遵循程序B.改变聚合温度将改变聚合时间和结构(Doyle et al。,2015)。降低聚合温度导致凝胶形成较慢。我们假设降低温度导致原纤维聚合的成核位点减少,因此原纤维延伸得更多,并在图2中观察到更大的孔径。
    7. 预计按照该协议,使用激光扫描共聚焦,旋转盘共聚焦(图2)或光片显微镜可以容易地观察到单个原纤维。


      图2. 4%标记胶原蛋白的荧光图像。 A.单一共焦切片(XY)和10%YZ投影(右)4%atto 488标记的3 mg / ml大鼠 - 尾胶原凝胶在21°C聚合。 B.图A中所示的胶原凝胶的3D渲染(X,Y,Z尺寸:分别为100×100×80μm)。 C.用EGFP-α-辅肌动蛋白(品红)转染的HT-1080纤维肉瘤癌细胞的高分辨率最大强度投影,其通过在16℃聚合的Atto 565标记的3mg / ml大鼠尾胶原(绿色)迁移。图像显示时间序列中的第一帧(左)和63 rd 帧(右)(Z每15微米,每隔30秒在Z上成像)。时间以分钟为单位。

笔记

虽然使用鼠尾胶原蛋白是一种使用3D水凝胶的简单方法,但它需要重复才能使凝胶变得一致。胶原蛋白I型3D凝胶高度依赖于1)胶原浓度,2)离子浓度,3)pH,当然还有4)温度。因为这些中的每一个都会影响聚合时间,这不可避免地导致ECM结构的差异,所以上述每个参数在实验之间是一致的是至关重要的。 pH值是迄今为止变化最大的,因为它需要进行测试和调整以中和每批胶原蛋白。建议每批应检查pH值,并且不要假设1 N NaOH的量总是相同。在整个方案中使用小等份的储备溶液以减少由于随时间发生的蒸发引起的浓度变化。
需要注意的其他关键问题是10x DMEM解决方案:由于盐度高,在4°C或冰上时总会出现沉淀物。研磨10倍DMEM溶液对于保持一致的离子浓度是重要的。

食谱

  1. 10x DMEM(50 ml)
    1. 将1份含酚红(Sigma-Aldrich)包装的粉末DMEM加入50ml蒸馏水中
    2. 用热(约50℃)搅拌混合物直至DMEM粉末进入溶液中
    3. 温暖时无菌过滤。制备4个10ml和20个0.5ml等分试样。 0.5ml大小的等分试样用于日常实验
    4. 将两种尺寸储存在-20°C直至使用。将热等分试样在37°C水浴中解冻,然后在冰上涡旋并冷却
    5. 可在-20°C下无限期保存,在4°C下保存1个月
  2. 10x重建缓冲液(100 ml)
    1. 2.2克碳酸氢钠(Sigma-Aldrich)
    2. 4.8克HEPES(Sigma-Aldrich) - 或 - 20毫升1M HEPES原液,最终0.2 M
    3. 蒸馏水可达100毫升
    4. 过滤灭菌并在-20°C下储存成类似于10x DMEM的等分试样
    5. 可在-20°C下无限期保存,在4°C下保存1个月
  3. 氢氧化钠溶液(NaOH),1N
    1. 0.5g NaOH颗粒(Sigma-Aldrich)
    2. 蒸馏水至12.5毫升
    3. 充分混合并过滤灭菌,分成500μl等分试样并无限期保存在-20°C
  4. 1 N HCl
    1ml HCl(37%或12M,Sigma-Aldrich)
    蒸馏水至12毫升
    过滤灭菌,分成500μl等分试样并无限期保存在-20°C
  5. 50 mM硼酸盐缓冲液(pH 9.0)
    1. 1.55g硼酸(粉末99.5%,Sigma-Aldrich)
    2. 加蒸馏水至400毫升
    3. 在搅拌的同时一次加入几个固体NaOH颗粒(Sigma-Aldrich)直至pH为~9.0
    4. 加蒸馏水至500毫升
    5. 使用0.2μm过滤器过滤消毒
    6. 在室温下储存长达1年
  6. DMSO中的5 mg / ml荧光NHS-酯染料(200μl)
    1. Atto-488 NHS-酯1mg(Sigma-Aldrich)
      注意:任何NHS-酯染料都是合适的,可以替代
    2. 将200μlDMSO(Sigma-Aldrich)加入染料管中并在旋转混合器上混合(用箔包裹)1小时
    3. 立即使用并以10-20μl等分试样储存多余的
    4. 储存在-80°C
  7. 氯化钠(NaCl),8%溶液
    1. 将4 g NaCl(Sigma-Aldrich)溶于500 ml蒸馏水中,充分混合
    2. 使用0.2μm过滤器过滤灭菌
  8. 50 mM Tris缓冲液(pH 7.4)
    5 ml 1 M Tris(KD Medical)
    加蒸馏水至100毫升
    使用0.2μm过滤器过滤消毒
    在室温下储存长达1年

致谢

我要感谢Joshua Collin,Gavrel Pacheco和Tomoko Ikeuchi的有益评论。这项工作得到了NIDCR校内研究部的支持。这项工作改编自以前的工作(Doyle,2016)。作者声明没有利益冲突。

参考

  1. Chandrakasan,G.,Torchia,D。A.和Piez,K。A.(1976)。 从大鼠尾腱和皮肤制备完整的单体胶原蛋白,并在溶液中制备非螺旋末端的结构。 J Biol Chem 251(19):6062-6067。
  2. Doyle,A.D.,Carvajal,N.,Jin,A.,Matsumoto,K。和Yamada,K.M。(2015)。 局部3D基质微环境通过收缩性依赖性粘连的时空动态调节细胞迁移。 em> Nat Commun 6:8720。
  3. Doyle,A。D.(2016)。 生成具有受控多种架构的3D胶原凝胶。 Curr P rotoc Cell Biol 72:10 20 11-10 20 16.
  4. Fitch,S。M.,Harkness,M。L.和Harkness,R。D.(1955)。 从组织中提取胶原蛋白。 自然 176(4473 ):163。
  5. Friedl,P.,Noble,P.B.,Walton,P.A.,Laird,D.W.,Chauvin,P.J。,Tabah,R.J.,Black,M。和Zanker,K.S。(1995)。 体外间充质和上皮癌外植体中协调细胞簇的迁移 。 Cancer Res 55(20):4557-4560。
  6. Gross,J.,Highberger,J。H.和Schmitt,F.O。(1955)。 通过中性盐溶液从结缔组织中提取胶原蛋白。 Proc Natl Acad Sci USA 41(1):1-7。
  7. Weiss,P。和Garber,B。(1952)。 间充质细胞的形状和运动作为培养基物理结构的功能:对定量形态的贡献。 Proc Natl Acad Sci USA 38(3):264-280。
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引用:Doyle, A. D. (2018). Fluorescent Labeling of Rat-tail Collagen for 3D Fluorescence Imaging. Bio-protocol 8(13): e2919. DOI: 10.21769/BioProtoc.2919.
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