Longitudinal Intravital Microscopy Using a Mammary Imaging Window with Replaceable Lid

All procedures described in this paper were performed in accordance with the guidelines of the IACUC and KU Leuven and were carried out within the context of approved project application P160/2020. Prior to executing this protocol, please review the analgesia strategy with the institutional veterinarian and administer pre-and post-operative analgesia as per institutional guidelines.

1. Preparation of the R.MIW (estimated timing: 2 h)

NOTE: For the construction of the R.MIW, it is advisable to find a local manufacturer who specializes in working with hard materials such as titanium, as this requires special tools and expertise. Workshops specializing in the design and manufacture of titanium prosthetics usually have the machinery and expertise to produce the R.MIW parts.

1. Place the lid of the R.MIW on a sterile surface with the outside of the lid facing upwards (and the arms of the lid sloping downwards; Figure 1A).
2. Apply cyanoacrylate adhesive glue around the entire rim of the lid and carefully place a 10 mm round glass coverslip on top of the lid.
   NOTE: Always prepare a spare lid with a glass coverslip as a backup during the surgical procedure.
3. Use a sterile wooden stick to position the glass coverslip and apply some force to push it down to ensure an air-tight seal between the glass coverslip and the frame of the lid. Make sure the holes in the arms of the lid are not covered up by the glass.
4. Disinfect the lids and the R.MIW by submerging in 80% ethanol for at least 30 min in a closed Petri dish.
   NOTE: Do not leave the lids longer than 1 h in 80% ethanol to avoid the dissolution of the cyanoacrylate adhesive glue.
5. Remove the excess cyanoacrylate adhesive glue from the glass coverslip by using a cotton tip soaked in acetone.
6. Store the lids and the R.MIW in a sterile environment until further use.
   NOTE: The R.MIW frame and lids can be kept in a sterile tube or bag for a maximum of 1 week. When resuming the experiment, always examine whether the glass coverslip is still properly attached to the lid prior to proceeding with the protocol.

2. Preparation for surgery (estimated timing: 15 min)

NOTE: For the R.MIW implantation procedure, female mice (>3.5 weeks old) of any strain can be used.

1. Maintain sterility by using sterile gloves, and sterilize all items that will come in contact with the mouse. Wash surgical tools with water and mild soap, air dry, wrap in aluminum foil with no exposed corners or edges, and sterilize using an autoclave for a cycle of 20 min at 120 °C prior to the surgery.
2. Prepare a sterile surgical platform and sterilize the surfaces with 80% ethanol.
   NOTE: To ensure sterility, the surgery can be performed in a flow cabinet.
3. Turn on the heating pad and cover the mat with a sterile drape. Layout the sterile instruments on the sterile drape and place the R.MIW and a minimum of two glass-covered lids in sterile phosphate-buffered saline (PBS) in a Petri dish (close the lid to avoid external contamination).
4. Anesthetize the mouse in an induction chamber using a 3% isoflurane/O₂ mixture. Ensure that the animal is relaxed enough to be easily manipulated and transferred to the nose mask without any struggle.
5. Transfer the mouse from the induction chamber to an anesthesia nose mask and reduce the isoflurane level to 1.5% – 2% isoflurane/O₂ mixture. Verify full anesthesia by performing a paw withdrawal test.
6. To perform the paw withdrawal test, firmly pinch the animal paw using fingernails or blunt end forceps. In the absence of any muscle reflex, consider the animal as unconscious and proceed for surgical preparation. Perform this test before animal manipulation and repeat regularly during the anesthetic and surgical procedures to confirm anesthesia.
7. Apply eye ointment to prevent corneal dehydration.
8. Shave the area around the 4^th mammary gland using a razor blade (Figure 2A). Use the 4^th nipple as a landmark. Remove loose hairs using sticky tape.
   NOTE: Alternatively, depilatory cream can be used to remove the hairs.
9. Disinfect the exposed skin with 80% ethanol or povidone-iodine and position the mouse in the sterile surgical area on its back with the snout into an anesthesia nose mask using 1.5% isoflurane/O₂ mixture.
10. Secure the front and hind limbs of the mouse using paper tape.
    NOTE: Leave the anesthetized and recovering mouse on the heating pad as much as possible, during and after surgery.
11. Use a precut sterile surgical drape or gauze to cover the mouse whilst leaving the surgical area exposed.

3. R.MIW implantation (estimated timing: 30 min)

1. Verify if the animal is adequately anesthetized by performing a paw withdrawal test. Gently lift the skin using a fine Graefe forceps and make a 10-15 mm diagonal incision using small spring scissors in the skin on top of the 4^th mammary fat pad using the 4^th nipple as an orientation point (Figure 2A). Make sure not to cut or damage the peritoneum or the mammary gland.
2. Define the region of interest (ROI) based on macroscopic features of the mammary tissue, including the location of the lymph node or a tumor lesion, and try to keep the ROI central with regard to the incision.
3. Separate the skin from the mammary fat pad and underlying tissue layers using blunt dissection of up to 5-8 mm all around the incision line to create a pocket to fit the R.MIW frame (Figure 2BI).
4. Pick up the R.MIW using sterile forceps and test whether the incision is large enough to insert the window. If needed, slightly enlarge the incision until the R.MIW fits in.
5. Remove the R.MIW and place a purse-string suture all around the incision using a 5-0 silk suture (braided, non-resorbable). Place the suture 1-2 mm from the edge of the incision from the outside to the inside of the skin, starting from the caudal end of the incision.
6. Move approximately 5 mm up along the incision and pass the suture thread through the skin from inside to outside. Repeat this procedure along the edge of the incision, thereby creating a circular suture consisting of 5-6 loops (Figure 2BII). The final exit of the suture thread should be located approximately 2-5 mm away from the first entrance.
   NOTE: Be careful not to place the suture too close to the edge of the incision, as this increases the risk of skin tear. Placing the suture too far from the edge of the incision increases the risk of infection between the excess skin and the groove of the R.MIW.
7. Fit the R.MIW inside the incision and use the Graefe forceps to carefully place the skin in the groove of the R.MIW. Make sure to leave the loops of the suture outside.
8. Pull on the loops of the purse-string suture and tighten the suture in the groove of the R.MIW by gently pulling on both ends of the suture (Figure 2BIII). Tie off with surgical knots and cut away the excess thread.
9. Apply petroleum jelly to the inner rim of the R.MIW using a toothpick, while making sure to avoid the underlying tissue. Gently push the window frame downwards with sterile forceps or two fingertips, and position the lid into the R.MIW frame (Figure 1B). This will avoid a volume of air in between the mammary gland tissue and the R.MIW lid.
10. Place the tips of thin forceps through the holes in the arms of the lid and tighten the lid by twisting the lid clockwise (approximately 5°) (Figure 1C and Figure 2BIV).
11. If some air remains after tightening the R.MIW lid, use a sterile insulin needle to remove the excess air. Introduce the needle through the skin into the cavity between the mammary tissue and the lid and slowly pull out the plunger, thereby creating a vacuum between the mammary gland tissue and the R.MIW lid.
12. Administer, post-surgery, analgesia such as buprenorphine (0.1 mg/kg diluted in sterile 0.9% NaCl) by subcutaneous injection. Repeat subcutaneous injection with buprenorphine at 8 h and 16 h after the surgery.
13. Put the mouse back into the cage to recover and monitor closely until it is fully awake, or proceed with step 4 for immediate imaging.
14. At this post-surgical stage, monitor the mice closely for signs of discomfort or complications based on vital parameters, such as respiration, reactivity, behavior, posture, and body weight. Monitor the skin surrounding the window carefully for signs of inflammation and necrosis. If no complications occur, the R.MIW will allow for repeated IVM sessions over several weeks post-implantation.

Figure 2: Overview of the surgical procedure and window implantation. (A) A diagonal incision is made near the 4^th nipple of a female mouse and the skin and underlying tissue are disconnected by blunt dissection. (B) The inguinal lymph node and shape of the fat pad can be used for the right orientation of the tissue (I). A purse-string suture is placed in the skin around the incision (II), and after fitting in the window frame, the suture is tightened in the groove to secure the window frame, and secured by surgical knots (III). The last step is the insertion and locking of the lid into the window frame (IV). Please click here to view a larger version of this figure.

4. Repeated intravital microscopy through the R.MIW

NOTE: The repeated IVM strategy described here was optimized for an inverted multiphoton confocal system equipped with a motorized stage and a dark climate chamber at 35.8 °C.

1. Anesthetize the mouse in an induction chamber using a 3% isoflurane/O₂ mixture. Verify loss of consciousness by performing a paw withdrawal test (described in step 2.6). Apply eye ointment to prevent corneal dehydration during the imaging session.
2. Inspect the condition of the R.MIW frame and lid, including the stability of the sutures or any potential damage to the lid. In case of damage to the lid, the lid can be replaced as described in step 5.
3. Transfer the mouse to a pre-heated (37 °C), custom-made imaging box (Figure 3A) and place the mouse with the head into the nose cone. The imaging box consists of a frame that fits within the automated stage of the microscope. The frame is designed to hold a custom-made inlay with a hole the diameter of the R.MIW (Figure 3A) .
4. An inlet with a nose cone is connected to the front of the box and attached to an isoflurane vaporizer station using 2%-3% isoflurane/O₂ mixture. An outlet is connected to an anesthetic scavenging unit to ensure circulation and to prevent accumulation of isoflurane in the box. The imaging box can be closed using a transparent lid.
5. Position the mouse on the inlay in such a way that the R.MIW falls in the hole of the imaging box inlay (Figure 3A). Apply parafilm and paper tape over the back of the mouse to stabilize the mouse and reduce tissue motion due to respiration.
6. Close the imaging box with the lid and insert the imaging box in the stage of the microscope.
7. Reduce the isoflurane dose gradually over the course of the imaging session to sustain a stable, but superficial anesthesia (typically between 1.5%-0.8% isoflurane/O₂ mixture).
8. Provide proper nutrition during imaging as described below.
   1. For imaging sessions <3 h, inject the mouse subcutaneously with 200-300 µL of sterile PBS for hydration.
   2. For imaging sessions >3 h, follow the steps outlined below.
      1. For long-term imaging, infuse the mouse with nutrients. For this, use a commercially available sterile infuse mixture containing amino acids and glucose. Place a flexible needle subcutaneously in the neck of the mouse and secure it with paper tape.
      2. Attach a 10 mL syringe with the prepared nutrient mix to a flexible silicon tube and push the nutrient mix through until it has reached the end of the tube.
      3. Connect the silicone tube to the flexible needle. Take the end of the tubing and place it through one of the holes of the imaging box (from inside to outside, leaving the needle attachment side in the box).
      4. Inject 50 µL of infuse solution every 30 min.
9. Secure the imaging box in the stage of the microscope and define the area of interest using the epifluorescence mode of the microscope.
10. Apply the desired microscope settings depending on the mouse model (Figure 3B). Vessel structures, collagen patterns (visualized by second harmonic generation), and other stable anatomical structures, including the inguinal lymph node, may be used as landmarks. Acquire images using an inverted multiphoton confocal microscope at a 12-bit depth, and a 25x water objective using a Z-step size ranging from 1.0 to 4.0 µm (total z-stack ranging from 150 to 800 µm). Apply the following standard settings: 1024 x 1024 format, 600 Hz scan speed, bidirectional mode.
11. Visualize collagen I fibers by performing second harmonic generation by using an excitation wavelength of 860 nm and detection at 425-435 nm. Excitation and detection wavelengths for the different fluorophores are indicated in Table 1.

Table 1. Please click here to download this Table.

12. Use the Spiral function in imaging software navigator mode to generate a large overview tilescan to create a reference map of the tissue (Figure 3B). Define the ROIs within the spiral overview, define the upper and lower Z planes to determine the Z stack. Press Start to obtain detailed three-dimensional (xyz) or four-dimensional Z-stacks (xyzt) of the tissue.
    NOTE: During the entire imaging experiment the mouse should be closely monitored. For long imaging sessions, a pulse oximeter and temperature probe are recommended. The breathing of the mouse should be regular and at the same pace. If the mouse starts to show signs of gasping, the amount of isoflurane should be reduced.
13. At the end of each imaging session, keep the mouse on a heating pad until fully awake. If the mouse shows signs of discomfort or reduced mobility, keep the cage for a longer time on the heating pad and provide nutrient gels instead of the regular chow.
    NOTE: In between imaging sessions, the mouse should be closely monitored for signs of discomfort such as decreased food and water consumption, rapid breathing, decreased movement, tremor, abnormal body posture, or unkept fur. In case of clear signs of discomfort, follow institutional guidelines regarding animal welfare and terminate the experiment when the humane endpoint is reached.
14. Repeat steps 4.1-4.12 for every repeated imaging session and use the overview tilescan to retrace the same position(s) over multiple time points (Figure 3B). Imaging frequency will depend on the research question and the timing of the studied process, and typically varies from imaging sessions twice a day to once per week.
15. At the end of the desired experimental period, euthanize the animals with an implanted R.MIW using deep anesthesia with isoflurane followed by cervical dislocation. If desired, dissect the organs of interest for further ex vivo analyses.
16. Alternatively, keep the animal alive for future in vivo analyses. In that case, remove the pouch suture holding the R.MIW by cutting it with spring scissors and carefully detach the window ring from the mouse skin using blunt forceps. Clean up the edges of the skin that previously held the window and proceed with a simple continuous suture (absorbable material, such as polyglycolic acid).
17. Once the skin is closed, tie the beginning and ending suture threads with two square knots (4 throws). To minimize tissue ischemia, the knots should be loose enough to allow blood flow at the skin edge. Disinfect the wound with povidone-iodine to prevent inflammation and to promote healing of the skin.

Figure 3: Longitudinal IVM workflow. (A) Schematic depiction of a typical multi-day IVM experiment. (B) Imaging of a tumorigenic region within the adult mammary gland. Prior to each imaging session, a low-resolution overview tilescan (top panel) can facilitate the identification of the region(s) of interest over subsequent imaging days. The collagen I pattern (second harmonic generation, magenta), tissue structure and patterns of differentially labeled cells with fluorescent proteins (depicted in green, yellow, and red) can be used to retrace the same tissue area. Scale bars represent 1 mm (overview tilescan) and 100 µm (bottom panels). (C) Schematic representation of the R26-Confetti reporter construct. Upon tamoxifen-induced Cre-recombination, the Confetti construct stochastically recombines, resulting in the expression of one of the four fluorophores (nGFP, YFP, RFP, or mCFP). Please click here to view a larger version of this figure.

5. Opening and closing the lid of the R.MIW in between imaging sessions

1. Maintain sterility by using sterile gloves, and sterilize all items that will come in contact with the mouse. Autoclave surgical tools prior to the surgery (see step 2 for details).
2. Turn on the heating pad and cover the mat with a sterile drape and anesthetize the mouse in an induction chamber using a 3% of isoflurane/O₂ mixture. Verify adequate anesthesia by performing a paw withdrawal reflex test.
3. Transfer the mouse from the induction chamber to an anesthesia nose mask and reduce the isoflurane level to 1.5%-2% isoflurane/O₂ mixture.
4. Place the tips of thin forceps through the holes in the arms of the lid and loosen the lid by twisting the lid anti-clockwise. If needed, apply some pre-heated sterile PBS (37 °C) on the window to facilitate the loosening of the lid.
5. Place the lid of the R.MIW in a sterile dish with sterile PBS until further use. Clean the coverglass of the lid with demi-water and subsequently 80% ethanol. Keep the lid in sterile PBS until further use.
6. Prevent the exposed tissue from drying out by adding some drops of sterile PBS to the tissue surface.
7. Apply or inject any substance or cells of interest on the exposed tissue, a maximum volume of 50 µL can be used. Wait a few minutes until the liquid is absorbed by the tissue. Alternatively, the tissue can be manipulated or repositioned using sterile, blunt forceps to optimize the imaging conditions for specific ROIs.
8. Apply petroleum jelly to the inner rim of the R.MIW, making sure to avoid the underlying tissue. Position the lid into the R.MIW frame as described in step 3.9, place the tips of thin forceps through the holes in the arms of the lid, and tighten the lid by twisting the lid clockwise (approximately 5°).