Longitudinal Intravital Imaging Through Clear Silicone Windows

Summary

An approach is here presented for long-term intravital imaging using optically clear, silicone windows that can be glued directly to the tissue/organ of interest and the skin. These windows are cheaper and more versatile than others currently used in the field, and the surgical insertion causes limited inflammation and distress to the animals.

Abstract

Intravital microscopy (IVM) enables visualization of cell movement, division, and death at single-cell resolution. IVM through surgically inserted imaging windows is particularly powerful because it allows longitudinal observation of the same tissue over days to weeks. Typical imaging windows comprise a glass coverslip in a biocompatible metal frame sutured to the mouse’s skin. These windows can interfere with the free movement of the mice, elicit a strong inflammatory response, and fail due to broken glass or torn sutures, any of which may necessitate euthanasia. To address these issues, windows for long-term abdominal organ and mammary gland imaging were developed from a thin film of polydimethylsiloxane (PDMS), an optically clear silicone polymer previously used for cranial imaging windows. These windows can be glued directly to the tissues, reducing the time needed for insertion. PDMS is flexible, contributing to its durability in mice over time—up to 35 days have been tested. Longitudinal imaging is imaging of the same tissue region during separate sessions. A stainless-steel grid was embedded within the windows to localize the same region, allowing the visualization of dynamic processes (like mammary gland involution) at the same locations, days apart. This silicone window also allowed monitoring of single disseminated cancer cells developing into micro-metastases over time. The silicone windows used in this study are simpler to insert than metal-framed glass windows and cause limited inflammation of the imaged tissues. Moreover, embedded grids allow for straightforward tracking of the same tissue region in repeated imaging sessions.

Introduction

Intravital microscopy (IVM), the imaging of tissues in anesthetized animals, offers insights into the dynamics of physiological and pathological events at cellular resolution in intact tissues. The applications of this technique vary widely, but IVM has been instrumental in the cancer biology field to help elucidate how cancer cells invade tissues and metastasize, interact with the surrounding microenvironment, and respond to drugs^1,2,3. In addition, IVM has been key to advancing the understanding of the complex mechanisms governing immune responses by providing insights complementary to ex vivo profiling approaches (e.g., flow cytometry). For instance, intravital imaging experiments have revealed details about immune functions as they relate to cell migration and cell-cell contact and have offered a platform to quantitate spatiotemporal dynamics in response to injury or infection^4,5,6,7. Many of these processes can also be studied through histological staining, but only IVM allows the tracking of dynamic changes. In fact, whereas a histological section offers a snapshot of the tissue at a given time, intravital imaging can track intercellular and subcellular events within the same tissue over time. In particular, progress in fluorescence labeling and the development of molecular reporters have allowed molecular events to be correlated with cellular behaviors, such as proliferation, death, motility, and interaction with other cells or the extracellular matrix. Most IVM techniques are based on fluorescence microscopy, which due to light scattering, makes imaging deeper tissues challenging. The tissue of interest, therefore, often needs to be surgically exposed with an often invasive and terminal procedure. Thus, depending on the organ site, the tissue can be imaged continuously for a period varying from a few to 40 h⁸. Alternatively, the surgical insertion of a permanent imaging window permits the imaging of the same tissue sequentially over a period of days to weeks^7,9.

The development of new imaging windows has been highlighted as a technological need to further improve intravital imaging approaches¹⁰. The prototypical intravital imaging window is a metal ring containing a glass coverslip secured to the skin with sutures¹¹. Interference with free movement, the accumulation of exudate, and damage to the glass coverslip are common problems seen with using such windows. Moreover, the prototypical window requires specialized production, and the surgical procedure can require extensive training. To address these issues, polydimethylsiloxane (PDMS), a silicone polymer, which has previously been used in cranial windows for long-term imaging in the brain¹², was adapted for use in abdominal organ and mammary gland imaging. Here, a detailed method for generating PDMS-based silicone windows is presented, including how to cast the window around a stainless-steel grid to provide landmarks for repeated imaging of the same tissue regions. Furthermore, a simple, stitch-free surgical procedure for inserting the window over abdominal organs or the mammary gland is described. This new approach overcomes some of the most common issues with currently used imaging windows and increases the accessibility of longitudinal intravital imaging.

Protocol

All procedures described were performed in accordance with the Cold Spring Harbor Laboratory Surgical Guidelines and had been approved by the Institutional Animal Care and Use Committee at Cold Spring Harbor Laboratory.

1. Casting the silicone window

1. Prepare the silicone polymer (PDMS) by mixing the base elastomer and curing agent in a 10:1 (v/v) ratio.
2. Cast a window by depositing a small quantity of PDMS on a sterile, smooth surface and adjust the volume-to-area ratio to the desired thickness.
   NOTE: Using 200 mg of polymer solution for a 22 mm diameter circle on the lid of a 24 well-plate lid results in a good compromise between window sturdiness and optical clarity.
3. Optional: To provide landmarks for repeated imaging of the same tissue regions, lightly press a stainless-steel grid into the silicone after the PDMS is on the desired casting surface.
4. To remove air bubbles, place the coated surface in a vacuum desiccator for 45 min to degas the polymer.
5. Cure the silicone windows in an oven at 80 °C for 45 min.
6. Score the polymer at the edges of the mold and gently peel the cured windows from the surface used for casting with forceps.
7. Before surgery, sterilize the windows by autoclaving.

2. Preparing the mouse for insertion of the silicone window

1. Anesthetize the mouse in an induction chamber using 4% (v/v) isoflurane. Move the mouse to a warming pad on the surgical table, place the mouse in the anesthesia nose mask, and lower the isoflurane concentration to 2% for maintenance throughout the surgery.
2. Check the depth of anesthesia by pinching the toes of the hind limbs. Do not proceed until the mouse is inactive and does not show toe pinch reflex response.
3. Apply ophthalmic lubricant to the eyes to keep them from drying and preventing trauma.
4. Administer pre-emptive analgesia (buprenorphine 0.05 mg/kg) subcutaneously.
5. Shave the surgical site and completely remove the hair with a hair removal cream.
6. Apply povidone-iodine solution (10% v/v) and ethanol (70% v/v) consecutively 3x to prevent the surgical site infection.

3. Inserting a ventral window for imaging in the liver; adaptable for other abdominal organs (Figure 1)

1. Place the mouse in the supine position.
2. Make a 10 mm incision starting 3 mm down from the xiphoid process using sterile scissors and forceps.
3. Remove a 1–1.5 cm² section of skin along the midline.
4. Use a second pair of sterile scissors and forceps to remove a section of the peritoneum slightly smaller than the section of skin.
5. Optional: To visualize a larger portion of the liver, use sterile cotton swabs moistened with sterile saline to push the liver down from the diaphragm revealing the falciform ligament connecting the liver to the diaphragm. Sever the ligament taking care not to cut the inferior vena cava.
6. Withdraw surgical adhesive with a 31 G syringe, apply a small quantity of it on the surface of the liver around the edges of the area to be imaged. This adhesive creates a circular seal, leaving the center area intact for imaging.
   CAUTION: Limit the adhesive to small droplets forming a circular pattern around the imaging area. Tissue immediately in contact with the adhesive cannot be imaged. It is not necessary to dry the tissue before applying the adhesive.
   NOTE: Any cyanoacrylate-based adhesives can be used successfully for this procedure. Surgical adhesive ensures sterility. For terminal procedures, all-purpose super glues yield good results.
7. Position the window using forceps and hold it firmly against the liver until the adhesive has dried (~2 min).
8. Fold the edges of the window under the peritoneum.
9. With a syringe, deposit a small quantity of surgical adhesive on the edges of the window that are now under the peritoneum. Push down on the peritoneum with forceps to secure it to the window.
10. Similarly, deposit a small quantity of surgical adhesive onto the peritoneum before pushing down on the skin with forceps to secure it to the peritoneum.
    NOTE: If performed correctly, a circular area of liver tissue should now be visible through the window.
11. Apply glue around the edges of the window to create a rim that will help prevent the skin from growing back over the window.
    CAUTION: If the procedure is carried out using sterile surgical techniques and the window is sterilized before insertion, sealing the wound with surgical adhesive is sufficient to avoid infections. However, failure to follow proper aseptic techniques during surgical insertion or imperfect closure can lead to overt or subclinical infection and drying out of the tissue.

4. Inserting a lateral window for imaging in the liver; compatible with the concurrent injection of cancer cells in the portal vein (Figure 2)

1. Place the mouse on the left lateral decubitus position
2. Make a 10 mm incision on the right flank 3 mm below the costal arch using sterile scissors and forceps.
3. Remove a 1 cm² section of skin.
4. Use a second pair of sterile scissors and forceps to remove a section of the peritoneum slightly smaller than the section of skin.
5. Withdraw surgical adhesive with a 31 G syringe, apply a small quantity of it on the surface of the liver around the edges of the area to be imaged.
   CAUTION: Limit the adhesive to small droplets forming a circular pattern around the imaging area. Tissue immediately in contact with the adhesive cannot be imaged.
   NOTE: Any cyanoacrylate-based adhesives can be used successfully for this procedure. Surgical adhesive ensures sterility. For terminal procedures, all-purpose super glues yield good results.
6. Position the window using forceps and hold it firmly against the liver until the adhesive has dried (~2 min).
7. Fold the edges of the window under the peritoneum.
8. With a syringe, deposit a small quantity of surgical adhesive on the edges of the window that are now under the peritoneum. Push down on the peritoneum with forceps to secure it to the window.
9. Similarly, deposit a small quantity of surgical adhesive onto the peritoneum before pushing down on the skin with forceps to secure it to the peritoneum.
   NOTE: If performed correctly, a circular area of liver tissue should now be visible through the window.
10. Apply glue around the edges of the window to create a rim that will help prevent the skin from growing back over the window.
11. If a portal vein injection is required:
    1. Place the mouse in the supine position.
    2. Make a 10 mm incision starting down from the xiphoid process in the skin.
    3. Using a second pair of sterile scissors and forceps, make a 10 mm incision in the peritoneum.
    4. Dip two cotton swabs in sterile saline.
    5. Use the cotton swabs to gently pull the intestine onto sterile gauze moistened with sterile saline, taking care not to twist or otherwise upset the orientation.
    6. Keep displacing the intestine from the abdominal cavity until the portal vein is visible on the right side of the abdomen.
    7. Insert a 31–33 G needle 5–7 mm caudal to the point of entry of the portal vein into the liver, making sure to proceed within the blood vessel and not through it.
    8. Slowly inject the cancer cells (resuspended in 100 μL of sterile PBS).
       NOTE: For this experiment amurine pancreatic cancer cell line (e.g., KPC-BL/6-1199) can be cultured in complete DMEM media and 1 x 10⁵ cells injected in 100 μL of sterile PBS.
    9. To prevent bleeding, immediately after withdrawing the needle, apply gentle pressure on the injection site with a small piece of surgical sponge for 1–2 min.
    10. After the pressure is released, monitor the site for 1–2 min to ensure hemostasis.
    11. Using moistened cotton swabs, return the intestine into the abdominal cavity following the physiological orientation of the organ.
    12. Using sterile forceps, insert the window immediately under the peritoneum, on the left side of the abdominal cavity of the mouse.
    13. Suture with 4-0 silk sutures and staple the midline incision with 7 mm stainless steel wound clips.
    14. Move the mouse to the left lateral decubitus position
    15. Make a 10 mm incision on the right flank 3 mm under the costal arch through the skin using sterile scissors and forceps.
    16. Remove a 1 cm² section of skin around the incision.
    17. Use a second pair of sterile scissors and forceps to remove a section of the peritoneum slightly smaller than the section of skin.
    18. Pull the window into place over the liver before gluing it in place, as described under steps 4.8–4.10.

5. Inserting the window for imaging in the pancreas (Figure 3)

1. Place the mouse on the right lateral decubitus position.
2. Make a 10 mm incision on the left flank 3 mm below the costal arch using sterile scissors and forceps.
3. Remove a 1 cm² section of skin around the incision.
4. Use a second pair of sterile scissors and forceps to remove a section of the peritoneum slightly smaller than the section of skin.
5. Using sterile cotton swabs soaked with sterile saline, gently pull on the spleen to visualize the pancreas. Proceed to reposition the pancreas with the moistened cotton swabs to make more surface area visible through the incision.
   NOTE: At this point, pancreatic cancer cells can be orthotopically injected into the pancreas if it is a part of the experimental design.
6. Withdraw surgical adhesive with a 31 G syringe, apply a small quantity of it on the surface of the pancreas around the edges of the area to be imaged.
   CAUTION: Limit the adhesive to small droplets forming a circular pattern around the imaging area. Tissue immediately in contact with the adhesive cannot be imaged.
7. Position the window using forceps and hold it firmly against the pancreas until the adhesive has dried (~2 min).
8. Fold the edges of the window under the peritoneum.
9. With a syringe, deposit a small quantity of surgical adhesive on the edges of the window that are now under the peritoneum. Push down on the peritoneum with forceps to secure it to the window.
10. Similarly, deposit a small quantity of surgical adhesive onto the peritoneum before pushing down on the skin with forceps to secure it to the peritoneum.
    NOTE: If performed correctly, a circular area of pancreas tissue should now be visible through the window.
11. Apply glue around the edges of the window to create a rim that will help prevent the skin from growing back over the window.

6. Inserting the window for imaging in the mammary gland (Figure 4)

1. Place the mouse on the supine position.
2. Make a 10 mm incision medial to one of the inguinal nipples using sterile scissors and forceps.
3. Remove a 0.5 cm² section of skin above the mammary gland.
4. Use a second pair of sterile scissors to carefully separate the mammary gland from the skin by spreading the scissors between the two surfaces to disrupt adherence.
   NOTE: To facilitate imaging of the inguinal mammary gland area, be careful to dissect a portion of skin above the mammary gland but superior to the hind leg, so the window does not limit the mobility of the mouse.
5. Withdraw surgical adhesive with a 31 G syringe, apply a small quantity of it on the surface of the mammary gland around the edges of the area to be imaged.
   CAUTION: Limit the adhesive to small droplets forming a circular pattern around the imaging area. Tissue immediately in contact with the adhesive cannot be imaged.
6. Position the window using forceps and hold it firmly against the mammary gland until the adhesive has dried (~2 min).
   NOTE: A circular area of mammary gland tissue should now be visible through the window if performed correctly.
7. Fold the edges of the window under the skin.
8. With a syringe, deposit a small quantity of surgical adhesive on the edges of the window that are now under the skin. Push down on the skin with forceps to secure the skin to the window.
9. Apply glue around the edges of the window to create a rim that will help prevent the skin from growing back over the window.

7. Post-surgical recovery

1. Place the mouse in a clean recovery cage with ample nesting material, ensuring that part of the cage is resting on a heating pad.
2. Monitor the mouse continuously until it is conscious and mobile.
3. If needed, provide an additional dose of analgesic 12 h after the pre-emptive dose.
   NOTE: Additional analgesia is generally unnecessary, but consult with a veterinarian if the mouse shows signs of distress (such as hunched back, unkempt fur, or lost interest in food). Monitor the mouse daily for the first 3 days after surgery for signs of infection or other adverse effects. Less than 2% of mice require veterinary attention or euthanasia, typically due to partial detachment of the window. It is important to note that for male mice with ventral liver imaging windows, fighting between mice can result in detachment of windows. This is avoided by housing the mice separately.

8. Imaging through the window

1. Anesthetize the mouse in an induction chamber using 4% (v/v) isoflurane.
2. Apply ophthalmic lubricant to the eyes to keep them from drying and preventing trauma.
3. Move the mouse from the induction chamber to the microscope stage. Place the mouse in the anesthesia nose mask, and lower the isoflurane concentration to approximately 1–1.5% for maintenance anesthesia throughout the imaging procedure.
4. Place a pressure pad sensor below the mouse to monitor breath rate and fix the mouse to the stage using soft surgical tape.
5. Insert a rectal thermometer to monitor body temperature throughout the imaging session.
6. Turn on the heated pad, monitoring the mouse closely to ensure that body temperature does not exceed 37 °C.  
7. Utilize software to monitor the breath rate of the mouse. The optimal rate is ~1 breath/second. Adjust anesthesia as required.
8. Before each imaging session, clean the window from any residual lens immersion medium and debris by gently wiping it with a cotton swab dipped in 70% ethanol (v/v).
9. When utilizing a water immersion lens, apply ultrasound gel to the window, avoiding bubbles.
   NOTE: Distilled water can also be used but might require reapplication during imaging.
10. To find the optimal depth for imaging, first, locate the grid and place it in focus.
11. Determine the approximate tissue imaging depth by setting the bottom of the grid to 0. This information is necessary to locate the corresponding z plane in subsequent imaging sessions.
12. To identify the same location over multiple imaging sessions, utilize the squares on the grid as a reference point. During imaging, note the orientation of the grid and location within the grid of each field of view imaged (e.g., 2^nd row from the top, 4^th square from the left).
13. During successive imaging sessions, use the grid to navigate back to specific grid areas - and thus imaging fields.
14. Following each imaging session, remove any residual lens immersion medium and debris by gently wiping the window with a cotton swab dipped in 70% ethanol (v/v).

Results

Intravital imaging through imaging windows can be used to observe, track, and quantify a wide array of cellular and molecular events at single-cell resolution over a period of hours to weeks. Ideal features for an imaging window include: a) limited impact on the well-being of the mouse and the physiology of the tissue; b) durability; c) simplicity of insertion; and d) clear landmarks for repeated imaging of the same region. The result is a versatile, inert silicone window that is easily produced and inserted and that can...

Discussion

Intravital imaging windows are important tools for directly visualizing physiological and pathological processes at cellular resolution as they unfold over time. The novel procedure described for casting and inserting flexible, silicone imaging windows in mice overcomes some of the most common issues with currently used imaging windows (exudate, breaking, and interference with normal mobility), provides additional safety for the mouse, and increases the accessibility of this technique.

The mos...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
3M Medipore Soft Cloth Surgical Tape	3M	70200770819
Silk suture 4-0 PERMA HAND BLACK 1 x 18" RB-2	Ethicon	N267H
ACTB-ECFP mice	Jackson Laboratory	22974
AEC Substrate Kit, Peroxidase (HRP), (3-amino-9-ethylcarbazole)	Vector Laboratories	SK-4200
Alcohol swabs	BD	326895
Anesthesia system	Molecular Imaging Products Co.
Acqknowledge software and sensors	BIOPAC	ACK100W, ACK100M, TSD110
Betadine spray	LORIS	109-08
c-fms-EGFP (MacGreen) mice	Jackson Laboratory	18549
C57BL/6J mice	Jackson Laboratory	664
CD45 Monoclonal Antibody (30-F11)	Invitrogen	14-0451-82
CD68 Antibody	Abcam	ab125212
Curity gauze sponges	Covidien
Donkey Anti-Goat IgG H&L (HRP)	Abcam	ab6885
Donkey Anti-Rabbit IgG H&L (HRP)	Abcam	ab97064
Donkey Anti-Rat IgG H&L (HRP)	Abcam	ab102182
Dow SYLGARD 184 Silicone Encapsulant Clear	Electron Microscopy Sciences	24236-10	Two-part, 10:1 mixing ratio
Round Cover Glass, 8mm Diameter, #1.5 Thickness	Electron Microscopy Sciences	72296-08
Ender-3 Pro 3D printer	Shenzhen Creality 3D Technology Co., LTD
Far Infrared Heated blanket	Kent Scientific	RT-0520
Fc Receptor Blocker	Innovex Biosciences	NB309
Fiji imaging processing package			https://imagej.net/software/fiji/
FluoroSpheres carboxylate, 0.04µm, yellow-green (505/515)	Invitrogen	F8795
Gating system:	BIOPAC Systems Inc.		The components together allow monitoring mouse vitals during imaging and gating image acquisition on mouse respiration. All were acquired from BIOPAC systems.
Acqknowledge software		ACK100W, ACK100M
Diff. Amp. Module, C Series		DA100C
Dual Gating Sys small animal		DTU200
MP160 for Windows - Analysis system		MP160WSW
MouseOx Plus 120V		MOX-120V;015000
Pressure Pad		TSD110
Gelfoam	Pfizer	9031508	Absorbable gelatin sponge
Hardened fine scissors	Fine Science Tools	14090-11	Two pairs; stainless steel, sharp-sharp tips, straight tip, 26 mm cutting edge, 11 cm length
Human/Mouse Myeloperoxidase/MPO Antibody	R&D Systems	AF3667
Hot bead sterilizer	Fine Science Tools	18000-45	Turn on approximately 30 min before use; sterilize tools at >200 °C for 30 s
Imaris	Bitplane	www.bitplane.com
Immersion medium Immersol W 2010	Zeiss	444969-0000-000
Insulin Syringes with BD Ultra-Fine needle 6mm x 31G 1 mL/cc	BD	324912
Isoflurane (Fluriso)	VetOne	502017
Lycopersicon Esculentum (Tomato) Lectin (LEL, TL), DyLight® 594	Vector Laboratories	DL-1177-1
LysM-eGFP mice	www.mmrrc.org	012039-MU
Micro dissecting forceps	Roboz	RS-5135	Serrated, slight curve, 0.8 mm tip width; 4" length
Micro dissecting forceps	Roboz	RS-5153	1 x 2 teeth, slight curve, 0.8 mm tip width, 4" length
MTS MiniBionix II 808	MTS Systems		Servohydraulic material testing machine
Neutrophil Elastase 680 FAST probe	PerkinElmer	NEV11169
Nitrogen	General Welding Supply Corp.
Oxygen	General Welding Supply Corp.
Polylactic acid filament	Hatchbox		1.75 mm diameter
ProLong Diamond Antifade Mountant	Invitrogen	P36970
Puralube ophthalmic ointment	Dechra	NDC17033-211-38
Reflex 7 wound clips	Roboz Surgical	RS-9255
Stainless steel grid	Fotofab		One grid is 0.200 inches in diameter, with a total of 52 individual grid squares that are 0.016 x 0.016 inches. There is 0.003 inches of space between each square.
Surface Treated SterileTissue Culture Plates	Fisher Scientific	FB012929	Lid used as curing surface for imaging windows
TriM Scope Multiphoton Microscope	LaVision BioTec		Imaging was done on an upright 2-photon microscope (Trimscope, LaVision BioTec) equipped with two Ti:Sapphire lasers (Mai Tai and InSight, Spectra-Physics) and an optical parametric oscillator. The following Longpass Dichroic Beamsplitters (Chroma) were used to direct the signal towards four photomultipler tubes: T560LP T665LPXXR T495lxpr
Vetbond	3M	70200742529
VWR micro cover glass	VWR	48404-453