Intravital Two-Photon Microscopy of the Transplanted Mouse Lung

Summary

Here, we present a protocol to intravitally image the transplanted mouse left lung using two-photon microscopy. This represents a valuable tool for studying cellular dynamics and interactions in real-time following murine lung transplantation.

Abstract

Complications after lung transplantation are largely related to the host immune system responding to the graft. Such immune responses are regulated by crosstalk between donor and recipient cells. A better understanding of these processes relies on the use of preclinical animal models and is aided by an ability to study intra-graft immune cell trafficking in real-time. Intravital two-photon microscopy can be used to image tissues and organs for depths up to several hundred microns with minimal photodamage, which affords a great advantage over single-photon confocal microscopy. Selective use of transgenic mice with promoter-specific fluorescent protein expression and/or adoptive transfer of fluorescent dye-labeled cells during intravital two-photon microscopy allows for the dynamic study of single cells within their physiologic environment. Our group has developed a technique to stabilize mouse lungs, which has enabled us to image cellular dynamics in naïve lungs and orthotopically transplanted pulmonary grafts. This technique allows for detailed assessment of cellular behavior within the vasculature and in the interstitium, as well as for examination of interactions between various cell populations. This procedure can be readily learned and adapted to study immune mechanisms that regulate inflammatory and tolerogenic responses after lung transplantation. It can also be expanded to the study of other pathogenic pulmonary conditions.

Introduction

Lung transplantation is the final option for many patients suffering from end-stage pulmonary disease; however, long-term survival after lung transplantation is poor compared to other solid organ transplants. Survival at 5 years is only ~60%-70%¹, compared to 80%-90% in hearts² and 85%-90% in kidneys³. Many complications after lung transplantation, such as primary graft dysfunction, antibody-mediated rejection, and chronic lung allograft dysfunction, are due to the host immune response to the allograft. For example, our group has shown that neutrophils are rapidly recruited into the lung allograft following transplant-induced ischemia-reperfusion injury and form dynamic clusters surrounding blood monocytes⁴. Crosstalk between donor and recipient cells is responsible for deleterious alloimmune responses^5,6,7, and the ability to study these dynamic cell interactions in a live animal model is invaluable.

Two-photon microscopy allows for high-resolution intravital imaging for depths up to several hundred microns with minimal photobleaching of tissues^8,9. It is used in a variety of tissues and anatomic sites, including the neocortex^10,11, skin^12,13, and kidney^14,15. More recently, it has been adapted to non-static organs such as the lung and heart^4,16,17. In this protocol, we describe a technique to image stabilized, ventilated, and perfused pulmonary grafts following murine orthotopic left lung transplantation. A key benefit of the transplant model is the ability to genetically manipulate the donor and recipient separately. Individual cell populations can be visualized with transgenic mouse strains with knock-in fluorescent protein expression, adoptive transfer of fluorescently-labeled cells⁵, or intravenous injection of fluorescently-labeled antibodies to bind cell-specific markers^4,16,17,18.

In order to stabilize the lung during imaging, this protocol involves gluing the lung to a coverglass, while other groups have described suction stabilization using a custom-made reversible vacuum device¹⁹. Our protocol has several advantages, including a larger area of imaging and relative ease of setup using commonly available materials in a microscopy lab (including coverglass and glue). Since this gluing technique constrains the lung at the upper surface, it is expected to decrease ventilatory motion and allow for deeper imaging. This intravital imaging technique allows for detailed observation of immune cell behavior and interactions in real-time, which contributes to the study of immune mechanisms that regulate inflammatory versus tolerogenic responses following lung transplantation.

Protocol

All animal handling procedures were conducted in compliance with the National Institutes of Health Care and Use of Laboratory Animals guidelines and approved by the Institutional Animal Care and Use Committee at Washington University School of Medicine.

1. Anesthesia and intubation

NOTE: Orthotopic mouse left lung transplant is performed, as previously described^20,21. Lungs from 20-25 g C57BL/6 (B6) mice are transplanted into sex- and age-matched B6 recipients. B6.LysM-GFP reporter mice are used as recipients for select experiments to visualize neutrophil infiltration into lung grafts. Recipient mice can be imaged immediately following a lung transplant.

1. Re-anesthetize mice with intraperitoneal administration of ketamine (80−100 mg/kg) and xylazine (8−10 mg/kg). Administer sustained-release buprenorphine (0.5-1.0 mg/kg) subcutaneously for additional pain control. Confirm adequate depth of anesthesia with a paw pinch.
   NOTE: If time between lung transplantation and planned intravital imaging is less than two hours, can redose anesthetic with 50% of initial dose of ketamine.
2. Remove the hair from the entire left chest using an electric clipper and wipe the chest to remove the excess clipped hairs.
3. Apply non-medicated ophthalmic ointment to the eyes of the mouse to prevent corneal drying.
4. Re-intubate the mouse orotracheally with a 20 G angiocatheter, as previously described²¹.
5. Ventilate with room air at a rate of 120 breaths/min and tidal volume of 0.5 cc.
6. Deliver 1% isofluorane endotracheally during the entire procedure.
7. To visualize blood vessels during imaging, inject 20 µL of 655 nm nontargeted quantum dots (q-dots) in 50 µL of phosphate-buffered saline intravenously at least 5 min prior to imaging.

2. Surgical preparation of left lung for imaging

1. Place the mouse in a right lateral decubitus position (see Figure 1A).
2. Disinfect the chest skin with a mixture of 0.75% iodine and 70% ethanol, three times.
3. Re-open the left thoracotomy (from mouse lung transplant) through the third intercostal space (see Figure 1A).
4. Remove a portion of the skin and soft tissue over the left thoracotomy site, measuring 1.5 cm x 1.5 cm (see Figure 1B).
5. Prior to rib resection, first clamp the ribs for ~10 s to thrombose any microvasculature to achieve hemostasis (see Figure 1C, D).
6. Resect portions of the 3^rd to 5^th ribs to create an imaging window large enough to extricate the lung (~0.8 cm craniocaudally and ~1.0 cm anteroposteriorly, Figure 1D, E).
7. Stop any additional bleeding from the rib edges with electrocautery.
   NOTE: It is critical to achieve hemostasis in the cut edge of the ribs to avoid excessive blood loss during the imaging period.
8. Place a small bump (4 cm x 4 cm gauze folded longitudinally 3 times) under the right chest (see Figure 1E).
9. Using cotton swabs, elevate the lung graft out of the chest and place the lower aspect of the lung onto a 1 cm x 3 cm strip of saline-soaked gauze (see Figure 1F).
   NOTE: This strip of saline-soaked gauze prevents sliding of the lung, maintains a moist environment for the lung, protects the lung from the sharp edges of the resected ribs, and separates the lung from heart motion. Once the circumferential imaging chamber is applied over the lung (Figure 1G, H, described in section 3), there should be minimal heat and fluid loss from the open thoracic cavity during the imaging period.

3. Imaging chamber preparation

NOTE: An imaging chamber is custom-built (see Figure 2A). This imaging chamber consists of a base plate and a top plate between which the mouse is placed and secured in place with spring-loaded bolts on either side (Figure 2B). The top plate contains a circular cutout measuring ~2 cm in diameter. A correspondingly sized black O ring is placed into this opening on the front side of the top plate, which will protect the objective lens (Figure 2C). A 24 mm x 50 mm coverglass is adhered to the back of the top plate using high vacuum grease, which will create a watertight seal. This coverglass will serve as the imaging window by adhering to the lung below and holding the imaging media (i.e., water) above. Ensure that no vacuum grease gets within the circular imaging window. The coverglass will be replaced for each new imaging experiment. The base plate is heated to 35-37 ˚C using a thermocouple temperature probe (Figure 2A).

1. Place the intubated mouse onto the base plate of the imaging chamber with the open left chest facing up (see Figure 1G, H). Secure the mouse with silk tape over the face, front paws, and tail (see Figure 1H and Figure 2A).
2. Apply glue to the bottom side of the cover glass attached to the top plate (grey circle in Figure 2C, bottom), leaving a 0.8-1.0 cm glue-free circle in the center.
   NOTE: There is a tradeoff between the size of the glue-free circle and the amount of motion artifact; thus, it is recommended that the circle does not exceed 1.0 cm in diameter.
3. Lower the top plate until the coverglass makes contact with the lung, with the glue touching the lung surface (see Figure 1G).
   NOTE: The left lung will be glued to the coverglass with a clean, glue-free circle in the middle, which is the area that will be imaged.
4. Hold the coverglass against the lung and inflate the lung for 1-2 s so that the area of glue adheres to the surrounding tissue. Accomplish lung inflation by occluding the outflow tubing from the mouse endotracheal tube to the ventilator.
   NOTE: Do not inflate the lung for longer than 1-2 seconds, as this may cause excessive barotrauma.
5. Back off the top plate slightly to reduce pressure on the lung tissue. Ensure that the area of the lung within the imaging window does not move with ventilation.
6. Secure both ends of the top plate by securing the thumb nut over each bolt (see Figure 2B).

4. Two-photon imaging

NOTE: A fixed-stage, upright microscope with a 20x or 25x >1.0 numerical aperture (NA) water immersion objective should be used for intravital microscopy. Below is the setup used in this study for a B6 to B6.LysM-GFP left lung transplant with 655-nm q-dot blood vessel-labeling. When applying this protocol, the setup of the microscope, lasers, and dichroic filters can be adapted based on the needs of the specific experiment and the fluorescent reporters used.

1. Place the entire imaging chamber with the intubated recipient mouse (B6 left lung transplanted into B6.LysM-GFP with 655-nm q-dots) onto the microscope stage (see Figure 2A).
2. Use dichroic filters of wavelengths 480 nm, 560 nm, and 635 nm, which will appropriately separate the signals from the GFP reporter, q-dots, and second harmonic generation (SHG) signal (445 nm) generated by collagen. These filters can be adapted based on the specific experiment.
   NOTE: The SHG signal generated by collagen demarcates dark crater-like structures, which represent alveolar airspaces (see white arrows in Figure 3).
3. Set the titanium-sapphire femtosecond pulsed laser to 890 nm (within the near-infrared range) for excitation. Use the lowest possible laser power to achieve a sufficient signal.
   NOTE: The laser settings should be adjusted for the specific experiment to maximize signal separation.
4. Apply ~1 mL of water to completely cover the coverglass on the top plate, which will be kept in place by the O-ring and the vacuum grease seal of the cover glass to the top plate (see Figure 1G and Figure 2C).
5. Use the 20x >1.0 NA water immersion objective on the microscope. Ensure that the objective lens is adapted based on the specific experiment.
   NOTE: Larger NA allows for more light collection, higher contrast, and more detailed images. Water immersion objectives are preferred as water or buffer is more compatible with biological tissues.
6. Lower the objective of the microscope into the water and focus on the surface of the lung.
7. Activate the base plate heater and maintain the temperature at 35-37 ˚C.
   NOTE: If there are concerns about the inability to maintain tissue temperature or adequate perfusion to the lung, two additional resisters can be attached to warm the top plate in addition to the base plate (see Supplementary Figure 1).
8. Acquire image stacks with acquisition software of choice.
   NOTE: Ensure that the room is completely dark during acquisition, which may involve the use of blackout curtains/blinds and strategic covering of all light sources.

5. Video acquisition

NOTE: The following parameters can be adapted based on the specific experiment. Steps 5.1-5.3 describe the specific parameters used for the B6 to B6.Lysm-GFP murine left lung transplant, which can be used as a reference guide.

1. Using a video-rate bidirectional scanner, set the x-y scan dimensions to 512 x 512 pixels.
2. Using an xyz stepper unit, acquire time-lapse recordings of a 21-step z-stack with 10-frame averaging per step. Each stack takes about 20 s to acquire, which includes the time required for frame-averaging, the stepper motor to move and check its position, and a 100 ms delay.
   NOTE: The bidirectional scanner on the microscope used in this study operates at a video rate of 30 frames/s. This may differ depending on the specific microscope setup.
3. Acquire 80 consecutive z-stacks to obtain time-lapse imaging of leukocyte migration within the tissue parenchyma. This takes about ~27 min at 20 s/stack.
   NOTE: The stack size, frame-averaging, laser power, and total length of the time-lapse recording should be minimized to prevent excessive laser exposure to the tissue. If fluorescent reporters used are bright (i.e., GFP) and laser power is low, then repeating a 21-step z-stack with a 10-frame average, 512 x 512-pixel scan, at 20 s intervals for ~27 min is well tolerated by the mouse. If longer periods of imaging are desired, the acquisition time per stack should be increased to keep the total laser exposure the same (i.e., 1 min per stack over ~1.5 h). These settings will need to be optimized based on the fluorescent reporters used.
4. Euthanize the mouse at the end of image acquisition. To euthanize, anesthetize the mouse with ketamine (80−100 mg/kg) and xylazine (8−10 mg/kg), followed by subsequent cervical dislocation. Typical imaging preparations will easily maintain mouse viability for up to 2 h.
   NOTE: To extend this period, additional fluid supplementation, anesthesia with isoflurane, and attention to temperature regulation may be required.
5. Complete video rendering with Imaris. Other imaging software (such as ImageJ) can also be used.
   NOTE: Images may be processed post-acquisition using contrast enhancement, smoothing, and channel arithmetics to remove channel crosstalk.

Results

After 1 h of cold ischemic storage at 4 ˚C, we orthotopically transplanted the left lung from a B6 mouse into a B6.LysM-GFP mouse⁴, and then intravital two-photon imaging was performed, as described above. We performed imaging at two time points post-transplant - 2 h (Figure 3A) and 24 h (Figure 3B). Blood vessels are labeled in red by the q-dots injected immediately prior to imaging. Additionally, we can visualize monocytes that hav...

Discussion

Two-photon excitation was first described in her doctoral thesis by Maria Göppert-Mayer in 1931, who later won the Nobel Prize in Physics for describing the nuclear shell structure^22,23. Traditional fluorescence microscopy relies on single-photon excitation, with excitation wavelengths that are shorter and higher energy than emission wavelengths. In contrast to single-photon microscopy, two-photon microscopy involves simultaneous excitation by two photons, e...

Materials

Name	Company	Catalog Number	Comments
0.75% povidone-iodine	Aplicare	NDC 52380-0126-2	For disinfectant
1-inch 20G IV catheter	Terumo	SROX2025CA	For endotracheal tube (ETT)
1-inch silk tape	Durapore	3M ID 7100057168	To secure mouse in position
20x water immersion long objective lens	Olympus	N20X-PFH
3M Vetbond glue	Medi-Vet.com	10872	To glue coverglass to lung
655 nm non-targeted quantum dots	ThermoFisher	Q21021MP	For labeling of blood vessels
70% ethanol	Sigma Aldrich	EX0281	For disinfectant
Argent High Temp Fine Tip Cautery Pen	McKesson	231
Black O ring (2 cm)	Hardware store	N/A	For custom-built imaging chamber
Bolt (2)	Hardware store	N/A	For custom-built imaging chamber
Brass thumb nut (2)	Hardware store	N/A	For custom-built imaging chamber
Buprenorphine 1.3 mg/mL	Fidelis Animal Health	NDC 86084-100-30	For pain control
Chameleon titanium-sapphire femtosecond pulsed laser	Coherent	N/A
Cover glass (24 mm x 50 mm)	Thomas Scientific	1202F63	For custom-built imaging chamber
Curved mosquito clamp (1)	Fine Science Tools	13009-12
Dual channel heater controller	Warner Instruments	TC-344B
Fine scissors (1)	Fine Science Tools	15040-11
Fixed-stage upright microscope	Olympus	BX51WI
Gauze (cut to 1 cm x 3 cm)	McKesson	476709	To place under left lung
High vacuum grease	Dow Corning	N/A	To adhere coverglass onto top plate
Isoflurane 1%	Sigma Aldrich	26675-46-7	For anesthesia
Ketamine hydrochloride 100 mg/mL	Vedco	NDC 50989-996-06	For anesthesia
Metal sheet (3 cm x 7 cm)	Hardware store	N/A	For custom-built imaging chamber
Pointed cotton-tipped applicators	Solon	56225	To manipulate lung and for blunt dissection
Power Pro Ultra clipper	Oster	078400-020-001
Puralube Vet eye ointment	Medi-Vet.com	11897	To prevent eye dessiccation
Small animal ventilator	Harvard Apparatus	55-0000
Straight forceps (1)	Fine Science Tools	91113-10
Three channel shutter driver	Uniblitz	VMM-D3	Resonant scanner
x.y.z optical stepper motor	Prior Scientific	OptiScan II
Xylazine 20 mg/mL	Akorn	NDC 59399-110-20	For pain control