Imaging the beating heart in the mouse using intravital microscopy techniques - PubMed (original) (raw)

Imaging the beating heart in the mouse using intravital microscopy techniques

Claudio Vinegoni et al. Nat Protoc. 2015 Nov.

Abstract

Real-time microscopic imaging of moving organs at single-cell resolution represents a major challenge in studying complex biology in living systems. Motion of the tissue from the cardiac and respiratory cycles severely limits intravital microscopy by compromising ultimate spatial and temporal imaging resolution. However, significant recent advances have enabled single-cell resolution imaging to be achieved in vivo. In this protocol, we describe experimental procedures for intravital microscopy based on a combination of thoracic surgery, tissue stabilizers and acquisition gating methods, which enable imaging at the single-cell level in the beating heart in the mouse. Setup of the model is typically completed in 1 h, which allows 2 h or more of continuous cardiac imaging. This protocol can be readily adapted for the imaging of other moving organs, and it will therefore broadly facilitate in vivo high-resolution microscopy studies.

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Conflict of interest statement

COMPETING FINANCIAL INTERESTS The authors declare no competing financial interests.

Figures

Figure 1

Figure 1

Schematics of the protocol. This protocol illustrates a methodology for cardiac intravital microscopy in the mouse. The protocol is divided into six different phases. Phase 1, from Steps 1–9, details the electronics and the microscope setup for imaging and data acquisition. Phase 2, from Steps 10–20, illustrates the necessary steps for anesthesia and animal monitoring during the entire duration of the protocol. Phase 3, from Steps 21–34, details the surgical procedure for cardiac intravital microscopy. Phase 4, from Steps 35–40, explains the correct procedure for stabilizer positioning. Phase 5, from Steps 41–45, gives the necessary steps for both microscopy imaging and data acquisition. Finally, Phase 6 from Steps 46–48 highlights data processing routines for final image reconstructions. The approximate cost for the hardware, software and all reagents that are necessary to perform the protocol is in the range of 650,000,650,000, 650,000,6,000 and $3,000 (in USD), respectively. All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of the Massachusetts General Hospital, and they are in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

Figure 2

Figure 2

Mechanical tissue stabilizers. Mechanical tissue stabilizers can guarantee reproducibility in the motion over a physiological cycle during intravital microscopy imaging. (a) 3D-printed stabilizer used in the current protocol. The presence of a grid bonded to the pericardium provides efficient suppression of motion artifacts for identification and tracking of individual cardiomyocytes. (b) Suction-based stabilizer. Copyright 2012, from ‘Motion compensation using a suctioning stabilizer for intravital microscopy’ (ref. 15) by Vinegoni, C. et al. Adapted by permission of Taylor & Francis LLC,

http://www.tandfonline.com

.

Figure 3

Figure 3

Scheme of principle for gated sequential segmented microscopy. The ‘Frame Active’ timing waveform from the PSU microscopy unit indicates the time at which images are collected when operating in sequential mode. Although a single image has a duration _T_S, the time between sequentially acquired images is slightly longer (_T_F) because of the reversal of the galvanometer mirror in the scanning unit. On the basis of the acquired ECG signal, single image segments are extracted from sequentially acquired images in correspondence with the cardiac time gating window _T_GW. Because cardiac-induced motion is minimal during diastole, _T_GW is chosen in coincidence with this phase. If a sequential shift is introduced appropriately, the isolated segments can be combined in sequence to give rise to a final motion artifact–free reconstruction. _T_CC corresponds to the time interval between two cardiac cycles. All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of the Massachusetts General Hospital, and they are in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

Figure 4

Figure 4

Cardiorespiratory gating. Timing diagram taking into consideration respiratory-induced motion components. The ventilator trigger signal and ECG are recorded and two time gating windows (cardiac _T_CGW and inspiration _T_R) are located at points of minimum displacement. _T_R is usually selected near the end of the inspiration or expiration phase. A new gating window, _T_SGW, equal to the intersection between the two, needs to be considered in order to isolate within all images all segments that are representative of the same volume.

Figure 5

Figure 5

Scheme of principle for prospective sequential segmented microscopy. Microscope acquisition operates at a frequency compatible with a physiologic heart rate. A pacemaker operates at a frequency that is slightly different from the microscope frame-acquisition frequency (_T_CC < _T_F). Corresponding to different phases of the cardiac cycle (here represented in different colors), it is possible to isolate specific gating windows. By combining all segments from sequentially acquired images, final motion artifact–free images can be reconstructed at all phases of the cardiac cycle. All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of the Massachusetts General Hospital, and they are in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

Figure 6

Figure 6

Overview of the essential equipment setup necessary to use the protocol. (a) Surgical tools. (b) Stereomicroscope. (c) Ventilator and intubation tilting work stand. (d) Laser-scanning microscope for intravital imaging. (e) A typical integrated imaging platform. (f) Image acquisition and data monitoring is displayed during an intravital imaging session.

Figure 7

Figure 7

Schematic of the cardiac intravital microscopy system for confocal and two-photon imaging. Timing waveforms from an animal ventilator and an ECG monitoring amplifier are recorded in combination with the ‘Frame Active’ signal from the microscopy PSU unit. When you are performing prospective sequential segmented microscopy, a triggered pacemaker device is also inserted (red wirings in the figure) to actively pace the heart at a frequency slightly different from the microscope acquisition frequency.

Figure 8

Figure 8

Basic assembly of the stabilizer from the 3D machined parts. (a) 3D rendering of the stabilizer. (b) Components necessary to assemble the stabilizer. (c) Glue can be used to guarantee a permanent assembly of the different components. (d) Assembled stabilizer. (e) View in scale of the distal stabilizer ring unit.

Figure 9

Figure 9

Highlights from two different phases of the protocol. Important steps during phase 2 (animal monitoring and anesthesia) and phase 3 (surgical preparation) of the protocol are highlighted. (a) Isoflurane gas anesthesia vaporizer and delivery chamber. (b) Insertion of the tail vein i.v. line. (c) Primary skin incision. (d) Cautery through the superficial muscle layer to reveal the ribs. (e) Separation of the fourth intercostal space to access the pleural space. (f) Open thoracotomy incision. (g) Exposed heart with the pericardium intact. (h) Pacemaker lead sutured to the ventricular surface. (i) Expanded view of the area indicated with a green box in h. All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of the Massachusetts General Hospital, and they are in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

Figure 10

Figure 10

Right lateral decubitus positioning. The drawing illustrates the mouse in the right lateral decubitus position with the left front leg stretched in the cranial direction (Step 22).

Figure 11

Figure 11

Pectoralis muscle. (a,b) Two views of the pectoralis muscle. The two views indicate how changing the illumination path and angle view with the surgical microscope can help to clearly identify the pectoralis muscle. P, pectoralis muscle. R, rib. IC, fourth intercostal space. Care needs to be taken to not injure the muscle if survival surgery is planned (Step 27). All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of the Massachusetts General Hospital, and they are in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

Figure 12

Figure 12

Positioning and intravital imaging of the mouse. (a) A surgical drape covers the animal during the procedure, except for the exposed thoracotomy. (b) The animal is placed under the microscope on a heated stage. (c) The stabilizer also sits on the stage, and it is positioned using a translation stage (d). Imaging is performed without the objective lens contacting the mouse or the stabilizer. (e) A low-magnification lens (2×) helps to identify the region of interest and to position the high-magnification objective at the center of the stabilizer. (f) After positioning, the high-magnification objective is immersed in water for imaging. All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of the Massachusetts General Hospital, and they are in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

Figure 13

Figure 13

Positioning of the heart stabilizer. Different phases during the positioning of the stabilizer. (a) Step 35. (b) Step 37. (c) Step 38. (d) Step 39. Water is placed in the stabilizer. (e) Magnified view corresponding to the red box in d. (f) Water fills the stabilizer. (g) Step 40. All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of the Massachusetts General Hospital, and they are in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

Figure 14

Figure 14

Intravital imaging of the structure and function in the beating heart at cellular resolution. (a,b) Sequential segmented microscopy with retrospective gating enables multichannel fluorescence confocal microscopy of the heart in diastole over a large field of view (a) and at single-cell resolution (b). Reproduced with permission from ref. , Nature Publishing Group. (c–f) Prospective gating allows motion artifact–free imaging of cardiomyocytes at all phases of the cardiac cycle. (c–e) Measurements of the contractile changes of a single cardiomyocyte are obtained at three distinct time points of the cardiac cycle (f). The colored symbols on the cardiac cycle (f) show the stage of the cardiac cycle at which the images shown in c–e were taken. From left to right, the symbols correspond to the images shown in c–e. Scale bars, (a) 200 μm; (b) 10 μm; (c–e) 20 μm. All animal procedures and protocols were approved by the Institutional Animal Care and Use Committee of the Massachusetts General Hospital, and they are in accordance with the NIH Guide for the Care and Use of Laboratory Animals.

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