Fibered Confocal Fluorescence Microscopy (Cell-viZio™) Facilitates Extended Imaging in the Field of Microcirculation
A Comparison with Intravital MicroscopyLaemmel E.a · Genet M.b · Le Goualher G.b · Perchant A.b · Le Gargasson J.-F.c · Vicaut E.a
aLaboratoire d’Etude de la Microcirculation, Université Paris 7, bMauna Kea Technologies, et cLaboratoire de Physiopathologie cellulaire et moléculaire de la Rétine, INSERM U592, Paris, France Corresponding Author
This study investigated the capability of fibered confocal fluorescence microscopy (FCFM) to provide in vivo microvascular observations. FCFM is specifically designed for in vivo in situ observation thanks to a probe composed of a fiber bundle and micro-optics having a diameter as small as 650 μm. In the first part of the study, we compared the main characteristics of FCFM with those of intravital fluorescence microscopy (IFM). A mouse cremaster preparation was used as a common basis to allow for imaging with both modalities. We discussed the feasibility of obtaining quantitative measurements usually provided by IFM in the context of FCFM: morphometry, capillary permeability, functional capillary density, vasoconstriction and dilation effects. In addition, the possibility to visualize fluorescent red blood cells or leukocytes was also evaluated. Phototoxicity issues and limitations of FCFM were also discussed. We showed that FCFM allows observations and measurements usually provided by IFM and that the real-time capability of the system, as well as the flexibility and small diameter of the optical probe enable micro-invasiveness and can extend imaging capabilities for in vivo in situ observations when compared to IFM.
Copyright © 2004 S. Karger AG, Basel
Since the early 1970s, in vivo measurements made by intravital fluorescence microscopy (IFM) have proven to be crucial for the understanding of the physiology and the pathophysiology of the microcirculation.
Several experimental models have been proposed for the intravital exploration of a large number of organs in animals. Most experimental setups involve a surgical procedure, which makes it possible to place the studied organ under the objective of a microscope. The extent and invasiveness of the surgical procedure obviously depends on the amount of space required to allow the positioning of a microscope objective in the vicinity of the studied microvascular field. Therefore, it appeared to us that the possibility to minimize the dimension of the optical device to be used for microvascular exploration might represent an important qualitative step for intravital microscopy.
A new device (the Cell-viZioTM) combining confocal fluorescence microscopy with fiber optics has been recently developed. This new technology, referred to as fibered confocal fluorescence microscopy (FCFM), replaces the objective of a microscope with a micro mini-optical probe having a diameter as small as 650 μm.
Application of this novel technology is presented here in the context of the experimental study of microcirculation. In the first part of this study, a preparation of cremaster muscle was used as a common basis for comparison of FCFM and IFM. A comparison of vessel diameters measured using both techniques is presented. We also explored the possibility to evaluate functional capillary densities, changes in permeability, vasoconstriction and dilation. In addition, we also evaluated the possibility to visualize fluorescent red blood cells (RBC) or leukocytes. Phototoxicity has also been evaluated.
The second part of the study discussed the means offered by extended imaging using FCFM compared to conventional microscopy. We showed that in vivo in situ observations could be performed thanks to minimally invasive surgical procedures. Images of organs with limited accessibility such as the kidney, liver, mesentery, and conjunctiva are presented.
Materials and Methods
A conventional fluorescence microscope (Leitz, Wetzlar, Germany) equipped with a ×20 objective (numerical aperture = 0.32) and ×10 oculars was used as baseline equipment for comparison with FCFM. Epifluorescence illumination was performed with a xenon lamp, a heat-protecting filter, and an excitation filter. Images were captured using an acquisition and analysis software (Scion Image 1.62) controlling a charge-coupled device camera (COHU, high performance CCD), whose images were digitized using a frame grabber (LG-3 CCIR; Scion; 768 × 512 pixels, 25 images/s). Total magnification from the tissue to the computer screen was ×750. Field of view (FOV) was 300 × 200 μm.
The Cell-viZioTM (Mauna Kea Technologies, France: www.maunakeatech.com) is an FCFM imaging system . FCFM is based on the principle of confocal microscopy, which is the ability to reject light from out-of-focus planes and provide a clear in-focus image of a thin section within the sample. This optical sectioning property is what makes the confocal microscope ideal for imaging thick diffusing biological samples. The FCFM is especially designed for in situ and in vivo imaging thanks to a probe in length and diameter compatible with minimally invasive surgical procedure. A FCFM system is composed of three main components (fig. 1): (i) a laser-based optoelectronics unit; (ii) a range of optical fiber mini-probes (composed of tens of thousands of optical fibers) which is the link between the scanning device and the micro-objective, and (iii) a software unit to control the system and manage the image data. In the following section, we describe each of these components.
Fig. 1. Schema of the FCFM technique.
A 488-nm laser source, compatible with fluorescent dyes usable in vivo, is scanned by two mirrors on the proximal surface of a fiber bundle. Real-time imaging is achieved using a 4-kHz oscillating mirror for horizontal line scanning and a galvanometric mirror for frame scanning. Resulting frame rate is 12 Hz. The laser sequentially injected in each fiber is focused on the tissue via a distal micro-objective. The focalized spot allows the excitation of the fluorophore preferentially at a specific depth in the tissue. The fluorescence collected by the micro-objective is then reinjected in the same fiber that was used for illumination. Therefore, the fluorescence follows the same path as the excitation light until the dichroic filter diverts it to the photodetector. The confocality of the system derives from the size of each fiber core. Indeed, a 2-μm core diameter serves as a pinhole for both excitation and collection, giving the probe its optical slicing capability. Finally, a raw image is reconstructed from each measurement (fig. 2, left image). Scanning amplitude and signal sampling frequency have been adjusted to perform a spatial oversampling of the fiber bundle, which is clearly visible on the raw image where one can visualize the individual fibers composing the bundle.
Fig. 2.a, b Image obtained from a phantom containing fluorescent microspheres (∅2 μm). c, d Image obtained from vessels of mouse cremaster muscle on an area with low fluorescence response. a, c FCFM raw data. b, d After image processing. Image FOV: 160 × 152 μm.
A dedicated image processing algorithm has been designed in order to suppress the inherent artifacts which occur when imaging through a bundle of fibers. For this purpose, a two-step calibration of the FCFM system has been designed:
First step: measurement of the intrinsic autofluorescence of each fiber. This is obtained by imaging a non-fluorescent sample.
Second step: measurement of each fiber transmission/collection rate. This second measurement is obtained by imaging a homogeneous sample of constant fluorescence.
When these two properties are known for each fiber composing the bundle, it becomes possible to reconstruct from the raw data (where signal may vary due to the difference in the physical properties of the fibers) measurements that are proportional to the fluorescence of the tissue sample, and therefore appropriate for quantitative measurements. Special care was also taken regarding the spatial calibration of the system. Given the spatial mapping of the fibers on the fiber bundle (Fujikura, Japan), distortions observed on raw data images were compensated by the image-processing software to generate undistorted images.
Once the calibration step is passed, the Cell-viZio is ready for use. As image processing runs in real time (12 images/s), the user is provided with images of enhanced contrast (the background signal produced by the fiber autofluorescence is suppressed), showing neither spatial distortion nor optical fiber signal modulation (i.e. the processing renders the fiber bundle equivalent to a conventional lens; fig. 2, right image).
Optical mini-probes are basically to the Cell-viZio what an objective is to a microscope. A range of flexible optical mini-probes is available with various optical parameters. Table 1 presents major characteristics of the three probes used in our study.
Table 1. Main characteristics of micro-optical probes
The first optical probe has a diameter of 1.8 mm (ref. HD-1800-2.5, fig. 3). This probe performs a 20-μm-thick optical section of the tissue at a depth of 80 μm with a lateral resolution of 2.5 μm. The FOV is 160 × 120 μm.
Fig. 3. Tip of a 1.8-mm diameter probe (HD probe).
The second optical mini-probe used for acquisition of the images presented here has a diameter of 1.5 mm (ref. S-1500-5.0). This probe provides images immediately below the surface of biological tissue, with a slice thickness of 15 μm and a lateral resolution of 5 μm. The FOV is 400 × 280 μm. Finally, the third probe used has a diameter of only 650 μm (ref. S-0650-5.0) and similar optical performances to that of the S-1500-5.0.
All probes can be decontaminated using first detergent (ampholysine 0.5%) and appropriate disinfectants.
A mouse cremaster preparation classically used by our team for IFM observations of the microcirculation under physiological and pathological conditions was used as a common basis for IFM and FCFM data acquisition (however, one should keep in mind that FCFM is designed for used in situ on unprepared organs).
Male Balb/c mice were anesthetized by intraperitoneal injection of 90 mg/kg sodium pentobarbital. A tracheotomy was performed to facilitate spontaneous breathing, and the right carotid was cannulated with a polyethylene catheter for the injection of fluorescent solution. The right cremaster muscle was surgically prepared for in vivo visualization, by a technique proposed by our group and described in detail elsewhere . Briefly, the muscle was detached from the scrotum and a transverse buttonhole slit about 5 mm long was made in the proximal part of the cremaster pouch. The testicle and epididymis, and the cremaster itself were then drawn out through the buttonhole. The small pedicle that attaches the cremaster to the testicle was tied up with two stitches and cut between them, to separate the cremaster completely from the testicle. To prepare the cremaster muscle for microscopy, a flexible extensible ovoid ring was made with metal wire (0.1 mm in diameter) and placed so that the cremaster acquired a racket shape. The ring was positioned so that the main cremaster artery was in the center of the upper surface of the racket. Throughout these procedures, the muscle was continuously bathed with warm saline solution. The muscle was continuously superfused at 2 ml·min–1 with modified Krebs-Henseleit solution containing (in mmol/l) 118 NaCl, 5.9 KCl, 1.25 CaCl2·2 H2O, 0.5 MgSO4·7 H2O, 28 NaHCO3 and 10 glucose. The temperature of the solution was fixed to 34.5°C in the cremaster chamber. By bubbling the solution with a 5% CO2-95% N2 gas mixture, we fixed the pH, PO2 and PCO2 of this solution in the muscle chamber at 7.43 ± 0.03, 25 ± 1.7 and 40 ± 1.0 mm Hg respectively.
After preparation of the cremaster, the stage was placed under the plate of the fluorescence microscope and the optical probe of the FCFM was fixed on the side of the objective of the microscope. FITC-albumin (500 mg/kg) or FITC-dextran 150 (75 mg/kg) was administered intra-arteriously after a stabilization period to allow visualization of the microsvascular network by FCFM. After recording a sequence with the microscope, the plate of the microscope was shifted to approach the optical mini-probe to the same field, and then finely adjusted on the z-axis in order to obtain the corresponding image.
First we calculated the mean relative error and the coefficient of variation of a series of measurements (n = 10) made on consecutive images acquired on calibrated phantoms (i.e. micrometric scale) with IFM and FCFM. In addition we measured the coefficients of variation of series of measurements made on consecutive images (n = 20) acquired in 10 arterioles. Then, in the cremaster muscle, we successively recorded a common area during 2 s at 15 frames/s with the IFM and 2 s at 12 frames/s with FCFM. For off-line analysis, the sequence was replayed frame by frame to obtain the best image, and 54 vascular diameters were measured from 6 mice with the analysis software (Scion Image 1.62). Representative images from IFM and FCFM are presented infigure 4. FCFM images were made using the HD probe, the high resolution of which is particularly adequate for such measurements. The data were analyzed using Bland-Altman plots as the appropriate statistical method for assessing agreement between two different methods where neither method yields the true values . The data are plotted as a scatter plot of the mean value of the two methods versus the differences between the two methods.
Fig. 4. Images obtained for the same vessels of the mouse cremaster muscle with IFM (a) and FCFM (b). White lines indicate locations for vessel diameter measurements. HD probe, FOV 160 × 120 μm.
Moreover, we checked the possibility to use FCFM for assessment of microvessel diameter changes by measuring the effect of topical administration of acetylcholine (10–5 M) in arterioles preconstricted by noradrenaline (10–7M).
After plasma labeling, the functional capillary density (FCD) can be estimated on the basis of the number of capillaries per micrometer, which are perfused with RBC at the time of observation. In muscle, FCD can be estimated by counting the number of capillaries intersected by a straight line. We evaluated the ability to obtain FCD evaluation with FCFM using S probe as we found its FOV (400 × 280 μm) appropriate for FCD estimations.
To quantify macromolecular leakage across vessels, 75 mg/kg body weight of fluorescein-isothiocyanate-labeled dextran (FITC-dextran) was injected intra-arteriously after a stabilization period. Effective microvascular permeability to dextran was based on the accumulation of fluorescent-labeled dextran in muscular tissue. Macromolecular leakage was induced with histamine (10–5M superfused for 5 min). Image acquisition was made with FCFM 5–20 min after FITC-dextran administration. The S probe was used as we found its FOV (400 × 280 μm) appropriate for such observations.
We used the modified procedure described by Butcher et al.  for labeling erythrocytes with FITC (Sigma, ref A2889). Erythrocytes were obtained from separate donor mice by heart puncture after administration of pentobarbital. Erythrocytes were separated by centrifugation (×1,000, 5 min) and washed four times in a phosphate-buffered saline adjusted to pH 7.4 (Sigma, ref P5119) containing 100 mg/l EDTA (Sigma, ref 43,178-8), resuspended in PBS (pH 8) containing FITC and incubated at 25°C for 2 h. The labeled erythrocytes were again flushed with the PBS and centrifuged several times to free the supernatant from fluorescent dye. After completing the preparation of cremaster, labeled erythrocytes were administered intra-arteriously (25 μl/10 g mouse), or the fluorescent probe of leukocyte (rhodamine 6G; Sigma, ref R4127) was administered topically at increasing doses. We recorded sequences only with FCFM (HD probe).
Phototoxicity was evaluated after administration of FITC-albumin (500 mg/kg) or FITC-dextran 150 (75 mg/kg). Acetylcholine- induced vasodilation in 32 arterioles preconstricted by noradrenaline was measured with FCFM before and after 2, 3 or 5 min of constant illumination.
The capability of the FCFM to provide an invasive access to new sites in situwas studied. After anesthesia, the appropriate fluorescent dye was injected and an incision was done to enable organ access. The operator used the optical mini-probe in a hand-held mode.
When a series of measurements was made on consecutive images of a micrometric scale we found a mean relative error of 0.7% for the IFM and 1.0% for the FCFM and coefficients of variation of 0.8 and 0.4% for the two devices, respectively.When the two devices were used for in vivo measurements, average coefficients of variation for a series of 20 measurements in 10 arterioles were 1.2% for the IFM and 1.3% for the FCFM.
Typical images obtained when comparing measurements made on the same vessels using the two devices are shown in figure 4. As shown in figure 5a, the correlation between measurements made from FCFM and IFM in 54 vessels was very high (R2 = 0.98). A Bland and Altman  analysis confirms the good agreement between measurements made by both methods (fig. 5b). A systematic bias has been identified over the entire range of diameters measured. This bias, characterized by the mean difference between the two measurements made on the same vessel, was equal to 1.2 μm. The repeatability coefficient, defined as two standard deviations of the differences between measurements made with the two modalities on the same vessel, was 7 μm.
Fig. 5.a Correlation between diameter measurements made with the microscope (Dmicro) and with FCFM (Dfcfm). b Bland and Altman graphical representation. ––– = mean difference; ― = mean difference ± repeatability coefficient.
In addition, as shown in figure 6, we checked that it was possible to study the different phases of microvessel diameter changes induced by vasomotor agents.
Fig. 6. FCFM images for assessment of microvessel diameter changes induced by vasomotor agents. a Baseline (vessel ∅ = 60 μm). b Noradrenaline constriction (vessel ∅ = 38 μm). c Acetylcholine vasodilation (vessel ∅ = 61 μm). HD probe image FOV 160 × 120 μm.
As shown in figure 7, we also checked that FCFM allowed the user to obtain images from which FCD can be easily estimated. For reasons discussed below, we did not make formal comparisons between the two devices for this parameter.
Fig. 7. FCFM images for FCD estimation. S probe, FOV 400 × 280 μm.
As shown in figure 8, FCFM also allowed the user to quantify changes in permeability using fluorescent macromolecules. Macromolecular leakage was estimated by computing the mean gray level in an extravascular area, and figure 9 represents corresponding quantitative fluorescence extravascular evolution with respect to time.
Fig. 8. Macromolecular leakage. From top to bottom, left to right images acquired with FCFM 5, 7, 9, 11, 15, 20 min after histamine suffusion, respectively. ROI from which extravascular fluorescence measurements were made on first image (S probe, FOV 400 × 280 μm).
Fig. 9. Extravascular fluorescence signal as a function of time. Baseline: t = 5 min.
We obtained easily exploitable images of circulating RBC with Cell-viZio (fig. 10a). However in vivo labelling of leukocytes using rhodamine 6G (Sigma, ref R4127) did not produce FCFM exploitable images at the concentrations usually used in IFM (17.5 μM) but only at higher (175 μM) and possibly toxic concentration (fig. 10b).
Fig. 10. Visualization of labelled cells in venules in cremaster muscle with FCFM (HD probe, FOV 160 × 120 μm). a Erythrocytes. b Leukocytes.
When the same microvascular field was illuminated for several minutes, we observed the formation of platelet aggregates which represents evidence of endothelial lesions due to phototoxicity. Such aggregates where not present if the probe was moved from one field to another. When we quantified the threshold for phototoxicity using FITC-dextran and measuring the loss of acetylcholine-induced relaxation in preconstricted arterioles, we found evidence of reduced endothelial-dependent vasodilation only for a constant illumination time ≥3 min (fig. 11).
Fig. 11. Effect of illumination time with FCFM on acetylcholine-induced vasodilation in arterioles preconstricted by noradrenaline.
Note, however, that when FITC-albumin rather than FITC-dextran was used, the phototoxicity was higher due to the loss of endothelial-dependent dilation after only 1 min of constant illumination of the same vessel.
Figures 12 and 13 illustrate the use of HD and S probes in various organs. In contrast to the extensive surgical preparation required for exploring these organs by IFM, the use of FCFM probes only required a minimal incision. In fact, the laparotomy can be as small as a few millimeters, and a simple contact with any organ in the peritoneal cavity allows the acquisition of the vascular network. All encapsulated organs can be readily imaged through their capsule, and in their native environment. S probes (fig. 13) provide large FOV that enable to characterize the vascular architecture of the organs. For example liver sinusoids or kidney arterioles circumventing the renal tubules are clearly visible, and a glomerulus could also be imaged. Using a buttonhole made in the ileum, it is also possible to use FCFM for visualization of the microcirculation of intestinal villi. Highlighted by their microvascularization, intestinal microvilli show their structure. HD probes (fig. 12) provide smaller FOV but a higher resolution, facilitating the acquisition of details on the vascular beds and to measure vessel diameters.
Fig. 12. Images obtained with HD probe on stomach (a), ear (b), mesentery (c), kidney (d) and conjunctiva (e). FOV 160 × 120 μm.
Fig. 13. Images obtained with S probe on the external wall of the stomach (a), mesentery (b), kidney (c), intestinal microvilli (d), external wall of the bladder (e), liver (f), conjunctiva (g), pancreas (h). White arrow represents 100 μm. FOV 400 × 280 μm.
Small animal imaging modalities can be characterized by their resolution and invasiveness. When macroscopic in vivo and in situ observations are required, several non-invasive imaging modalities are adapted to small animals: computed tomography scan, magnetic resonance imaging, positron emission tomography and ultrasound. These devices provide images with a spatial resolution of 50 μm (MRI, CT and ultrasound) to 1–2 mm (positron emission tomography) . Other modalities have been adapted to in vivo microscopic imaging, such as conventional fluorescent microscopy, fast confocal microscopy, multiphotonic imaging or optical coherence tomography (OCT). For this second set of devices, spatial resolution varies from 0.5 μm for confocal multiphoton to 20 μm for OCT. This gain in resolution comes with experimental setups involving surgical procedures: such technologies are therefore invasive when used for in situ observations. However, those microscopy techniques are being adapted to minimally invasive approaches by using a fibered probe instead of an objective. Two-photon microscopy through a single fiber with distal scanning is under investigation [6, 7]. A fibered microprobe has been designed for endoscopic OCT acquisition . Various works on confocal microscopy potentially lead to in situ acquisition through a fiber bundle [9, 10] possibly using digital micro-device scanning . Using a single fiber, a distal micro-electro-mechanical systems scanning [12, 13] or a piezoelectric scanning  is under development. All these techniques aim at non-invasive, or minimally invasive (e.g. through a buttonhole) in vivo and in situ anatomical or functional imaging on small animals. However, most of these solutions are still limited by either the dimension of the optical probes (> 5 mm) [7, 12, 14] or the lack of real-time capabilities (0.5 Hz) . The FCFM system offers a compromise between optical-probe dimension (diameter as small as 650 μm), real-time capabilities (12 Hz) and spatial resolution (2–5 μm). Another difference of the FCFM system when compared to classical IFM is confocality. FCFM confocality enables optical sectioning with an axial resolution of 20 μm at about 80 μm below the tissue surface when HD probe is used. In some cases, this feature can simplify the observation by avoiding superimposition of vessels.
Recently, an orthogonal polarization spectral imaging technique (OPS) has also been proposed as a tool for exploring the microcirculation with reduced dimensions in comparison with IFM. This device has also been previously evaluated by our group . When compared with IFM the quality of images using OPS and FCFM were very close. However, even if the OPS probe is smaller than the optics of a microscope, it remains considerably larger than the FCFM probes. In comparison with the FCFM, OPS makes it possible to obtain microvascular observations without the application of a fluorescent dye [15, 16, 17]. This aspect can reduce toxicity but also limit some applications which can take advantage of fluorescence. Indeed, using fluorescence technique combined with confocality, visualization is not limited to organs which can be transilluminated. However, it should be stressed that the depth allowed by confocality in FCFM is limited to 80–100 μm. Taking advantage of the flexibility and small diameter of the probe, it becomes possible for some organs to avoid surgery by using natural ways  and for the others to minimize invasive surgery.
In the present study we addressed the ability of FCFM to provide in vivo microvascular observations and to be used for quantitative measurements most commonly made with IFM. The results of vessel diameter comparisons showed a high correlation coefficient between the measurements. The Bland and Altman graphical representation confirmed this result. Note that the 7-μm repeatability coefficient given by Bland and Altman could be partially due to the difference in resolution of the two devices (i.e. 1 μm for IFM and 2.5–5 μm for FCFM) and in addition it could be an overestimation since the images were not recorded simultaneously with the two techniques (it took between 30 s and 1 min for the operator to switch from the IFM to the FCFM acquisition). This means that the difference between the diameter measurements seen between the two techniques could be partially explained by spontaneous changes in diameter, which happen in a microvascular network. We found a systematic bias equal to 1.2 μm in the vessel diameter measurements, which could be at least partly due to the confocality of the FCFM. Moreover, the magnitude of the bias is inferior to the resolution of the FCFM. In addition, we found that the intrinsic variability of both devices, which were estimated from repeated measurements performed on calibrated phantoms or in in vivo arterioles, were very similar.
When using FCFM with the two probes that were tested it was possible to evaluate the density of perfused capillaries as is currently performed with IFM. Two reasons led us to avoid quantitative comparisons of the devices for this parameter. First, when no easily recognizable vessel pattern was available in a region of interest, it was almost impossible for the operator to switch from one modality to the other and to ensure that the same area was observed because of the difference in the FOV (about 300 × 200 μm for the IFM and 160 × 120 μm for the FCFM when the HD probe was used) and because of differences in image orientation. Secondly, due to the confocality of FCFM, FCD was performed in a fine optical section of 20 μm thickness while IFM would integrate capillaries from a much larger axial section.
The present study also allowed us to identify some limits of the present version of the FCFM. One limitation was the phototoxicity resulting in a deterioration of the vascular reactivity when the same site was continuously illuminated for 3 min or more. In addition, due to the sensitivity of the probes, the use of FCFM for leukocyte studies has been found to be very limited at the present time (this limitation might also apply to the use of FCFM for visualization of other fluorescent probes in vivo). In fact the sensitivity of the probe is related to the numerical aperture which depends on the diameter of the probe. The higher the numerical aperture is, the larger the probe diameter. There is obviously a trade-off between the reduced dimensions of the probe and its sensitivity. This point is under improvement with the application of innovative optical designs. Another improvement concerns the measurement RBC velocities, for which a software package is under investigation .
In conclusion, with the reduced dimensions of its optical mini-probes and its ability to image intact organs in their native environment, FCFM appears as an interesting tool in the study of the microcirculation. This is particularly true for organs with limited accessibility. Limited invasiveness, ease of use, and preservation of the physiology of the organ of interest are three benefits of FCFM which may facilitate the follow-up of animals during longitudinal studies. In addition, one well-known difficulty when studying the microcirculation with IFM is that responses from microvascular networks to similar pharmacological, physiological or physiopathological stimuli can be largely different from one organ to the other. Thanks to the diameter of the optical probes considered here, FCFM imaging may also be used as an adjunct to the IFM, thus enabling simultaneous observations of microvascular networks on several organs on the same animal. All sorts of combinations between different organs are possible as well as the use of more than one FCFM device at the same time.
The use of FCFM in the clinical situation has not been evaluated in the present study. Note, however, that FCFM can be used with fluorescein which is non toxic and commonly used for retinal angiography. Thus it can be considered that FCFM has a potential for clinical applications that should be explored in particular in the field of microcirculation.
Eric Vicaut, MD, PhD
Laboratoire d’Etude de la Microcirculation
10 avenue de Verdun
FR–75010 Paris (France)
Tel./Fax +33 1 40 05 49 73, E-Mail email@example.com
Received: June 24, 2004
Accepted: July 25, 2004
Published online: September 30, 2004
Number of Print Pages : 12
Number of Figures : 13, Number of Tables : 1, Number of References : 18
Journal of Vascular Research (Incorporating International Journal of Microcirculation)
Vol. 41, No. 5, Year 2004 (Cover Date: September-October 2004)
Journal Editor: U. Pohl, Munich; G.A. Meininger, College Station, Tex.
ISSN: 1018–1172 (print), 1423–0135 (Online)
For additional information: http://www.karger.com/jvr