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J Appl Physiol 102: 1235-1242, 2007. First published November 16, 2006; doi:10.1152/japplphysiol.00740.2006
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Technical and physiological background of plasma volume measurement with indocyanine green: a clarification of misunderstandings

Matthias Jacob,1 Peter Conzen,1 Udilo Finsterer,1 Alexander Krafft,2 Bernhard F. Becker,3 and Markus Rehm1

1Klinik für Anaesthesiologie and 3Physiologisches Institut, Vegetative Physiologie, Ludwig-Maximilians-Universität, Munich, Germany; and 2Klinik für Geburtshilfe, Universitätsspital Zürich, Zurich, Switzerland

Submitted 3 July 2006 ; accepted in final form 10 November 2006

ABSTRACT

The indocyanine green (ICG) dilution technique (DT) is frequently used for plasma volume (PV) measurement. However, because of inadequate knowledge about the properties of this dye, lack of accuracy has been attributed to the method. The aim of this report is to provide physiological background information about the ICG-DT to avoid some profound misunderstandings. When performing tracer dilution, one has to consider the tracer's distribution space before interpreting the result. For ICG, the distribution space is the total PV, i.e., circulating + noncirculating PV, fixed within the endothelial glycocalyx. The distribution space of red blood cells and large molecules, in contrast, is only the circulating part of PV. Therefore, it is erroneous to compare directly PV derived from different tracer dilution methods. The transcapillary escape rate of ICG should not relevantly influence measured PV if the method is performed properly, i.e., if a short time window of measurement is subjected to monoexponential extrapolation. A major problem of PV measurement in general is that the target itself is very inconstant. Thus, checking for constancy of ICG-DT with two consecutive measurements is unreliable. Nevertheless, the ICG-DT is a useful tool for determining PV, provided it is well understood by the investigator to enable correct interpretation of the results.

distribution space; plasma protein; endothelial glycocalyx; endothelial surface layer; albumin

EARLY EFFORTS TO MEASURE blood volume (BV) by dilution techniques (DT) were marked by an intensive search for a suitable tracer. In 1920, a systematic survey of ~60 dyes led to the use of "Evans blue" as a label for the plasma protein albumin (8). Evans blue, which was introduced into clinical use in the 1930s (12, 13), has been the standard plasma marker for a long time, despite its extended half-life (t = 100 min). Also radioiodinated serum albumin (t = 60 days), which is used to measure plasma volume (PV), and radiochromium-labeled red blood cells, which are used to measure red cell volume (RCV), became commonly used tracers; they are considered the "gold standard" for assessment of these two components of BV (19). However, the disadvantages of radioactive contamination and dye accumulation, especially in repeated measurements, are obvious. In addition, several studies revealed a possible mutagenic potential for Evans blue (25, 31). In 1968, indocyanine green (ICG) (3), a relatively unstable dye with an initial distribution volume comparable to that of Evans blue (16) and labeled albumin (28), was used for the first time to determine "blood volume." Further clinical applications are measurement of cardiac output (10) and liver function (6). Additionally, ICG serves as a stain for capillary microscopy, especially in ophthalmology (38). The risk profile is excellent: anaphylactic reactions have been reported only very sporadically (49).

In contrast to the assumptions of the first reporters, ICG, similar to Evans blue, is a plasma marker; therefore, whole BV cannot reliably be determined simply from ICG dilution. After this fact was recognized, ICG was frequently used as a tracer for PV measurements (14, 15, 22, 39–42).

Tracer dilution techniques for measuring PV are generally based on intravascular injection of a bolus of the tracer followed by measurement of the concentration in the patient's plasma after complete mixture (Fick's principle). The distribution space can be calculated from this concentration and the amount of dye injected, and the investigator normally assumes that this space represents PV. However, several reports questioned the accuracy of the ICG-DT for several reasons (20, 23, 29). This report aims to summarize the knowledge about the ICG-DT, including the physiological background, to forestall profound misunderstandings of a well-established method for direct measurement of total PV.

TRACER

ICG, a water-soluble and relatively unstable 755-kDa tricarbocyanine dye, rapidly binds to plasma proteins, mainly albumin, after injection. Complete mixture with the plasma can normally be expected 2 min after venous application (14, 15, 22, 39–42, 47, 48). It is rapidly and exclusively eliminated via the liver, by secretion into the bile, and t is only 2.5–3 min. ICG was found to be a safe, accurate, and precise tracer for PV measurements (14, 39, 47). Bound to plasma proteins, the peak extinction of ICG lies at 800 nm (Fig. 1). This wavelength also represents the "isobestic" point of hemoglobin, i.e., the wavelength at which the extinction of the oxygenated form equals that of the deoxygenated one. At 900 nm, ICG has no measurable extinction, in contrast to hemoglobin and other constituents of the plasma.


Figure 1
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  		Fig. 1. Light absorption spectra of indocyanine green (ICG), oxygenated hemoglobin (HbO2), and deoxyhemoglobin (Hb).

 
PRINCIPLE

Determining the concentration of any tracer before complete mixture with the plasma is pointless. However, the sought-after concentration at injection time (t0), but after complete mixture of the dye, never really exists. It is a theoretical value due to the overlapping processes of elimination and mixing, i.e., the time window required to allow complete mixture of the dye with the patient's plasma. To obtain the theoretical concentration at t0, one needs to determine the kinetics of early dye clearance. For this, it is necessary 1) to take several samples at reproducible times after complete mixture, 2) to measure the ICG concentrations in these samples, and 3) to extrapolate back to the theoretical concentration at t0.

EXTRAPOLATION TECHNIQUES AND TIME WINDOWS

At least during the first 5 min after recirculation, ICG elimination seems to consistently follow a monoexponential kinetic (48). However, beyond minute 5, the elimination kinetic inconstantly changes toward a biexponential, or even polyexponential, mode (24, 45, 48). To avoid overestimation of the distribution space of ICG (DSICG), the samples used for monoexponential extrapolation should only be taken up to minute 5. Otherwise, a "curve-fitting technique" is needed for computation of the theoretical concentration at t0 (48) from a biexponential elimination kinetic. For clinical use, the monoexponential extrapolation from values obtained during the first 5 min appears to be the better choice for several reasons: 1) it requires a shorter period of steady state of BV (important during surgery), 2) it provides PV almost "online" (see Whole blood method), 3) the investigator is not obliged to perform "black box" processing of the data, 4) immediate reinfusion prevents net blood loss (see Whole blood method), and 5) lower ICG doses (0.25 vs. 1 mg ICG per kilogram body wt) are required, accelerating recirculation.

DETERMINING ICG CONCENTRATION

ICG concentration is determined by measuring the extinction specifically caused by ICG. Therefore, it is necessary to subtract the basal extinction at 900 nm (no ICG extinction) from the extinction at 800 nm (peak ICG extinction) in each sample (Fig. 1). In venous blood, the oxygenation of hemoglobin is not constant; therefore, for measurement of the specific extinction of ICG in whole blood samples, we need arterial, fully O2-saturated blood to obtain a constant basal value at 900 nm. Before determination of unknown ICG concentrations, the system must be calibrated by measurement of two samples containing known ICG concentrations. This leads to the calibration equation (Fig. 2). Because ICG does not enter red blood cells but, rather, rapidly binds to plasma proteins after injection, we have to consider this only partial "dilution" of the dye in a whole blood sample (see Whole blood method) when calculating DSICG.


Figure 2
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  		Fig. 2. Calibration of the system before ICG injection. Respective specific extinction, resulting from 2 artificially generated dye concentrations in the patient's blood (whole blood method) or plasma (plasma method), leads to the calibration equation.

 
Plasma method.   For the plasma method (14, 15, 20, 29), only one single venous cannula, inserted into a cubital vein, is needed. After calibration of the system (measurement of the specific extinction in two 5-ml samples of the patient's plasma containing ICG at 2 artificially generated concentrations), the dye (0.25 mg ICG/kg body wt) is injected into the patient's circulation at t0. To prevent contamination of the venous catheter [mainly, the cone during connection and disconnection of the dye syringe (Fig. 3) ] and, consequently, contamination of the future blood samples with dye, even though the catheter is flushed with saline after injection, we use a second three-way valve (Fig. 4). This valve is removed immediately after the dye is injected and the system is flushed, i.e., before the first timed sample is withdrawn.


Figure 3
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  		Fig. 3. Schematic illustration of dye contamination of the cone of the venous catheter after disconnection of the dye-containing syringe, even though the catheter was flushed with saline.

 

Figure 4
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  		Fig. 4. Practical prevention of dye contamination of blood samples. A: when measurements are not being performed, the venous catheter is closed by a 3-way valve. B: immediately before injection, an additional 3-way valve is connected. C: syringe containing the dye is connected, and the dye is injected at time 0 (t0). D and E: crystalloid (10 ml) is immediately flushed through the connected system, consisting of 1 proximal and 1 distal 3-way valve in line with the venous catheter. F: potentially contaminated distal valve is disconnected and ejected. Proximal, clean valve remains and closes the venous catheter, as shown in A.

 
Starting exactly 2 min after t0, blood samples (2.5–3 ml each, taking ~3 s) are withdrawn every 20 s, up to the end of minute 5 (10 samples in all), with each withdrawal starting ~1.5 s before and ending ~1.5 s after the intended point of time. The catheter is not flushed between the two samples; rather, a few drops of blood are allowed to drip out immediately before the next sample is withdrawn (Fig. 5).


Figure 5
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  		Fig. 5. Blood withdrawal via the plasma method: placement of 16- to 17-gauge venous cannula (cubital vein), withdrawal of 3 ml/3 s in each sample, and no flushing between samples, rather, passive outflow of some drops of blood to allow fresh, venous blood to enter the catheter.

 
After separation of the red blood cells from the plasma by centrifugation, the specific extinction is determined in every sample. Specific extinction at t0 is derived by monoexponential extrapolation of the light absorption curve back to t0 (Fig. 6). This absorption value is incorporated into the calibration equation and leads to the theoretical plasma concentration of the dye at t0 (CP0). DSICG can then be calculated as follows: DSICG = D x CP0–1, where D is the amount of dye injected.


Figure 6
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  		Fig. 6. Theoretical extinction at t0, derived from monoexponential extrapolation of extinction values before the end of minute 5, leads to theoretical concentration of ICG at t0 when introduced into the calibration equation.

 
Menth-Meier et al. (28) demonstrated that it is appropriate to perform peripheral dye injection for PV measurement by ICG-DT. However, others advised induction of hyperemia before dye injection to accelerate recirculation (14, 28). Additionally, in preliminary, unpublished experiments (n = 10), we established that injection into one peripheral catheter and withdrawal of the timed samples from this and, simultaneously, an additional catheter on the other arm led to the same PV: difference = 11 ± 17 (SD) ml (P > 0.05, Student's paired t-test).

The "plasma method" is a scientific method that can be performed without causing discomfort to an awake volunteer. Because the samples must be processed as described above, it delivers DSICG at ~30 min after dye injection. Because ~40 ml of blood loss occurs per measurement, repeated measurements should be avoided to prevent our target, DSICG, from being relevantly influenced by the method.

Whole blood method.   In addition to a (central) venous cannula for tracer injection, an arterial line is needed for the whole blood method (22, 39–42). As explained above, at 800 nm, the oxygenation of hemoglobin is not relevant: the extinction is the same (isobestic point of hemoglobin). In contrast, extinction at 900 nm, leading to the basal value (see above), must be obtained with equally oxygenated hemoglobin in all samples.

As in the plasma method, immediately before the injection of the tracer, a two-point calibration, consisting of measurement of the specific extinction of two arterial blood samples containing two known, but distinct, ICG concentrations (Fig. 2), must be performed. Then the tracer (0.25 mg ICG/kg body wt) is injected into the (central) venous cannula at t0. For measurement of ICG concentration, blood is continuously drawn via the arterial catheter through a cuvette by means of a calibrated pump (20 ml/min). The blood is slightly heparinized and immediately reinfused. The cuvette is attached to the spectrophotometer, and as many timed measurements as possible are taken. We prefer taking measurements every 10 s between minutes 2 and 5 (22, 40–42).

The specific extinction at t0 is derived by monoexponential extrapolation of the light absorption curve resulting from the respective measuring time points (Fig. 6). If this absorption value is introduced into the calibration equation, the theoretical whole blood concentration of the dye at t0 (CB0) is obtained. CP0 is calculated as follows: CP0 = CB0 x (1 – HctLV)–1, where HctLV represents the large vessel hematocrit determined in blood samples withdrawn from the arterial line. DSICG is then calculated as follows: DSICG = D x CP0–1.

The whole blood method is much more convenient for the investigator than the plasma method and provides the plasma volume online, which can be important in the perioperative setting. Additionally, because of reinfusion of the samples, this method saves blood and, therefore, is ideal for intraoperative repeated measurements without impact on the target.

FROM DSICG TO PV: PHYSIOLOGICAL BACKGROUND OF A TECHNICAL METHOD

As explained above, when the ICG-DT is correctly performed, the result is DSICG, nothing else. The point is to correctly interpret this value to establish the "real" PV, and this is obviously a serious problem for some investigators (20, 23, 29): PV derived via ICG-DT was, e.g., "checked for correctness" by calculating RCV from PV and HctLV and expecting this RCV derived from ICG (RCVICG) to be constant in the absence of any bleeding or transfusion if the ICG-DT was consistent. In other cases, PV derived from ICG-DT (PVICG) was expected, as a proof for consistency of the method, to be constant before and after an intervention (e.g., induction of general anesthesia or volume loading). Such an assumption is invalid, however (see below). The transcapillary escape rate of albumin was also blamed to relevantly influence the results. However, central physiological properties of the microvasculature must be taken into consideration.

The fcell phenomenon.   There is an important, but inconstant, difference between HctLV and whole body hematocrit (HctWB), derived from measuring PV with the ICG-DT and RCV with fluorescein-labeled red blood cells. The quotient HctWB x HctLV–1, occasionally termed the "fcell ratio" in the literature, ranges from 0.73 to 1.10 (44). In some clinical trials, the fcell ratio was assumed to be 0.89–0.91 and was used as a correction factor to derive BV from PV or RCV and HctLV (5). However, the fcell ratio is not constant during many clinical interventions, e.g., volume loading (22, 40). Therefore, use of no factor or the same correction factor before and after such an intervention can lead to erroneous calculations of BV. This is illustrated in Table 1 by the "RCVICG," calculated without correction for fcell before and after volume loading. The calculation is based on HctLV and PV obtained using the ICG-DT (PVICG). As readily apparent, the reason for a seeming "decrease" in RCVICG without any actual loss of red blood cells is an impressive increase in the fcell ratio. Consequently, it would be erroneous to equate this apparent, not directly measured, RCVICG value with the real RCV derived from dilution of fluorescein-labeled red blood cells (RCVFl) (33). As a result, the premise that RCVICG must remain constant if red blood cells are not removed or lost (29) is not justified. Whether an intervention such as induction of general anesthesia, which is connected with vasodilatation and other hemodynamic responses, can also lead to a change in fcell ratio is not known. The fcell ratio does not seem to be systematically affected by sex, age, or systemic diseases. Corresponding work on animals (1, 27) and healthy (30, 46) and diseased humans (11, 52) confirmed only the well-known high variability mentioned above.


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  		Table 1. Measured and calculated PV and RCV before and after volume loading with hetastarch

 
What can be the reason for this variability in fcell, especially during interventions? A careful look at the individual distribution space of the different tracers used to determine PV offers a likely explanation. Recent intravital microscopic images revealed that there are actually three compartments of BV: 1) RCV, 2) a circulating part of PV (PVcirc), and 3) a noncirculating part of PV, which is bound by the endothelial glycocalyx (EG) (54). The first images of the EG showed only a thin (20–30 nm) layer, because conventional fixation techniques almost completely destroy the endothelial surface layer (ESL) before electron microscopy. In the past 15 years, however, indirect and direct approaches demonstrated that, at least in the small vessels, there is a 0.4- to 1.2-µm-thick ESL containing noncirculating plasma, including plasma proteins (Fig. 7). This exclusion zone for flowing red blood cells is in dynamic equilibrium with the flowing plasma (37). Investigations using fluorescence videomicroscopy of skin capillaries showed that ICG enters the ESL within 1 min (2, 4).


Figure 7
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  		Fig. 7. Electron-microscopic view of a capillary (left, lower magnification) and a part of the vessel wall (right, higher magnification). For staining of the endothelial glycocalyx, a modification of a method described by Vogel et al. (55), based on in situ stabilization of the glycocalyx by intracoronary application of a fixative containing La3+ and glutaraldehyde, was used (10, 26). TS, tissue space; VL, vascular lumen; EG, endothelial glycocalyx; PVcirc, circulating part of plasma volume; PVnoncirc, noncirculating part of plasma volume, representing plasma proteins fixed within the endothelial glycocalyx; PVtotal, total plasma volume (PVcirc + PVnoncirc).

 
Using the red blood cells as a tracer to quantify the circulating part of the plasma, we proceeded to compare PVICG with PVcirc, as derived from RCVFl and HctLV. Because PVICG represents the whole body PV, the difference was presumed to represent the volume of the whole ESL of the circulatory system. The total ESL determined in such a manner was 732, 717, and 431 ml in three separate experiments (22, 40). These findings were occasionally criticized as being unrealistic, since only 10% of whole body blood is found in systemic capillaries (23, 29). However, the ESL should not be restricted to capillaries. Although it was demonstrated in small vessels that this layer leads to a considerable decrease in hematocrit with respect to HctLV (37, 54), larger blood vessels are also lined by an ESL, possibly with an even larger dimension. If it is assumed that ~350 m2 is the total area of the endothelial interface with blood (37) and ~1 µm is the average thickness of the ESL (17, 54), a total ESL volume of ~350 ml can be expected. Because 1 µm is the assumed average height of the ESL in small vessels and it is likely that the whole body ESL is considerably thicker, our estimation introduced above seems quite reasonable. However, as demonstrated in Table 1, volume loading caused a dramatic decrease in the calculated noncirculating PV, presumably because of a decrease in ESL thickness. This, rather than a lack of consistency of the ICG-DT, is the reason for the apparent decrease in "RCVICG" in these patients.

If PV is the sought-after measure, one has to consider which part, the circulating or the total PV, is being stained (Fig. 7). Therefore, the properties of the ESL must be considered. It is an exclusion zone for large neutral and anionic molecules such as ≥70-kDa dextrans, but it is highly permeable for and, consequently, part of the distribution space of small neutral free dyes (e.g., 0.4-kDa rhodamine), plasma proteins such as albumin (53), and, consequently, tracers such as ICG and Evans blue, which bind rapidly to those plasma proteins. This fact underlines the statement of Tschaikowsky and co-workers (51) that "the smaller the indicator molecule, the bigger the [measured] plasma volume." However, it is probably not a faster vascular clearance of small molecules during the measurements that causes a "falsely too high" PV. The more likely reason for the finding of Tschaikowsky and co-workers is that, by using large molecules or particles that do not enter the ESL for PV measurement (e.g., stained red blood cells), only the circulating part of PV can be determined, which is lower than the total PV determined with tracers entering the ESL quickly (53). It may be expected that other large molecules, such as hydroxyethyl starch and large dextrans, do not enter the ESL within the few minutes taken for PV measurement and, therefore, should exhibit a distribution space more like stained red blood cells (Fig. 8) (53). Indeed, it has been demonstrated that there is no significant difference between the PV determined by using the tracer "10% hydroxyethyl starch 200,000/0.5" and that by using labeled red blood cells (51). Since the result should represent PVcirc directly, no fcell correction factor is necessary to calculate RCV from this PV and HctLV (26, 51). On the other hand, if it is assumed that the distribution space of albumin is PVcirc and the ESL, it is not at all surprising that the PV determined with radioactively labeled albumin is higher than the PV determined by red blood cells (50). Consequently, this phenomenon cannot be taken as a sign that the ICG-DT lacks precision.


Figure 8
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  		Fig. 8. Schematic drawing of distribution space (DS) of artificial colloids (e.g., hydroxyethyl starch or dextrans) and natural plasma proteins (e.g., albumin) at the vessel wall within the first minutes after injection, i.e., within the time window required for plasma volume measurement.

 
Transcapillary leakage of albumin.   Distrust in the ability of the ICG-DT to determine PV was also based on an indisputable transcapillary leakage of albumin (20, 23, 29). This exists even in normal subjects. Fleck et al. (9) reported a "transcapillary escape rate" of albumin of ~5%/h in healthy volunteers that can increase to 15%/h in the case of septic shock. On the basis of these values, "escape" of ICG attached to albumin molecules from blood vessels within our period of interest for PV measurements (0–5 min after injection of ICG) is low, normally ~0.4%, and rises to 1.25% under the worst clinical conditions (9). If it is assumed that this loss represents an increase in distribution volume, PV would be overestimated in a young man [180 cm height, 75 kg body wt, normal PV = 3,050 ml (36)] by only 12 ml, which would rise to 38 ml in the worst case. Obviously, this is not a relevant methodological pitfall for the ICG-DT.

Additionally, it is likely that there is no increased initial distribution volume of ICG due to the transcapillary escape rate together with albumin. Escape forced by attachment to albumin, even under the conditions of an increased capillary leak [e.g., caused by an increase of the capillary surface area (29)], should only add to the elimination rate normally dictated by the liver (18, 35). If the capillary leak of albumin does not change markedly within the 5 min of the PV measurement, the escape of ICG after the initial, rapid binding to plasma proteins (7, 34) and the initial distribution of the labeled proteins within the intravascular space will proceed at a monoexponentially constant rate. Consequently, monoexponential extrapolation, also under the conditions of a raised transcapillary escape rate, should lead to a quite correct estimation of the theoretical concentration of ICG at t0, although the elimination curve appears more steep in the monoexponential presentation.

Intraindividual inconstancy of PV.   PV is not a fixed individual quantity but, rather, varies due to physiological and pathophysiological stimuli. We found PV to be quite unstable shortly after induction of anesthesia or after an infusion therapy and surgical procedure (39). Changes in PV have been reported under the influence of different anesthetic drugs (32). As a result, a change in PVICG after an intervention such as induction of anesthesia is no legitimate basis for concluding that the ICG-DT is principally inaccurate (29).

Validation of the method.   Validation and testing of retest reliability are serious problems if the target of a method is intraindividually inconstant and each tracer has an individual distribution space. Nevertheless, the ICG-DT has been validated successfully. Comparison of two PV values derived from two different tracer dilutional methods is possible, if the respective tracers have a comparable distribution space. Because ICG rapidly binds to plasma proteins, validation of its distribution space using the distribution space of these proteins is justified. Validation with the reference method, radioiodinated albumin, was performed in 10 healthy volunteers and 21 patients after elective open-heart surgery, with excellent results even for peripheral ICG injection (28). In vitro and in vivo validation was performed by another group; however, they used a questionable reference tracer (radiochromium-labeled red blood cells) (5). This principal error in reasoning logically should not influence the in vitro results, inasmuch as there is no EG dividing PV into two compartments. The authors "saved" their mean in vivo results by entering a correction factor, namely, an assumed fcell ratio. In a comparison of ICG with Evans blue in 7 dogs and 16 patients, Haneda and Horiuchi (16) found a correlation coefficient of 0.98 between these two dyes. Although they erroneously assumed that they were measuring "total circulating blood volume," this misconception, inasmuch as the assumption was made for both markers, did not influence the information concerning validity of the ICG-DT.

Repeatability has been demonstrated for the plasma method and the whole blood method (14). A prerequisite is a steady state between the two measurements that are to be compared. Indeed, for assessment of validity or repeatability, at best, general anesthesia and full relaxation, or, at least, supine position for an extended period of time of the awake volunteer, should be implemented.

In conclusion, the ICG-DT can be a precise tool for measuring PV if it is completely understood by the investigator. In particular, one needs to recognize that the result obtained from this technique is, due to the tracer's distribution space, the total PV. Since this comprises the ESL, in addition to the circulating PV, it is larger than the distribution space of the red blood cells and large molecules. Consequently, direct comparisons with other methods to evaluate the accuracy of the ICG-DT are not applicable in most cases. In combination with direct measurement of RCV, the technique affords impressive insights concerning the integrity of the blood-tissue barrier. Besides routinely measuring total PV, the real target of infusion therapy during major surgery, relation of the total volume of the ESL to special (patho)physiological conditions or special interventions will be an important application of the ICG-DT of the future.

FOOTNOTES


Address for reprint requests and other correspondence: M. Jacob, Klinik für Anästhesiologie, Ludwig-Maximilians-Universität München, Nussbaumstr. 20, D-80336 Munich, Germany (e-mail: matthias.jacob{at}med.uni-muenchen.de)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

REFERENCES

  1. Barbee RW, Perry BD, Re RN, Murgo JP. Microsphere and dilution techniques for the determination of blood flows and volumes in conscious mice. Am J Physiol Regul Integr Comp Physiol 263: R728–R733, 1992.[Abstract/Free Full Text]
  2. Bollinger A, Saesseli B, Hoffmann U, Franzeck UK. Intravital detection of skin capillary aneurysms by videomicroscopy with indocyanine green in patients with progressive systemic sclerosis and related disorders. Circulation 83: 546–551, 1991.
  3. Bradley EC, Barr JW. Determination of blood volume using indocyanine green (Cardio-Green) dye. Life Sci 7: 1001–1007, 1968.[CrossRef][ISI][Medline]
  4. Brulisauer M, Bollinger A. Measurement of different human microvascular dimensions by combination of videomicroscopy with Na-fluorescein (NaF) and indocyanine green (ICG) in normals and patients with systemic sclerosis. Int J Microcirc Clin Exp 10: 21–31, 1991.[ISI][Medline]
  5. Busse MW, Zisowsky S, Henschen S, Panning B, Piepenbrock S. Plasma volume estimation using indocyanine green. A single intravenous injection method. Anaesthesia 48: 41–43, 1993.[ISI][Medline]
  6. Caesar J, Shaldon S, Chiandussi L, Guevara L, Sherlock S. The use of indocyanine green in the measurement of hepatic blood flow and as a test of hepatic function. Clin Sci 21: 43–57, 1961.[ISI][Medline]
  7. Cherrick GR, Stein SW, Leevy CM, Davidson CS. Indocyanine green: observations on its physical properties, plasma decay, and hepatic extraction. J Clin Invest 39: 592–600, 1960.[ISI][Medline]
  8. Dawson AB, Evans HM, Whipple GH. Behavior of large series of dyes introduced into the circulating blood. Am J Physiol 51: 232–256, 1920.[Free Full Text]
  9. Fleck A, Raines G, Hawker F, Trotter J, Wallace PI, Ledingham IM, Calman KC. Increased vascular permeability: a major cause of hypoalbuminaemia in disease and injury. Lancet 8432: 781–784, 1985.
  10. Fox IJ, Brooker LGS, Heseltine DW, Essex HE, Wood EH. A tricarbocyanine dye for continuous recording of dilution curves in whole blood independent of variations in blood oxygen saturation. Proc Staff Meet Mayo Clin 32: 478–485, 1957.
  11. Geers EA, Koomans HA, Boer P, Dorhout Mees EJ. Plasma and blood volumes in patients with nephrotic syndrome. Nephron 38: 170–173, 1984.[ISI][Medline]
  12. Gibson JG, Evans WA. Clinical studies of the blood volume: application of a method employing the azo dye Evans blue and the spectrophotometer. J Clin Invest 16: 301–316, 1937.[ISI][Medline]
  13. Gregersen MI, Gibson JG, Tead EA. Plasma volume determination with dyes; errors in colorimetry; use of the blue dye T-1824. Am J Physiol 133: 54, 1935.
  14. Haller M, Akbulut C, Brechtelsbauer H, Fett W, Briegel J, Finsterer U, Peter K. Determination of plasma volume with indocyanine green in man. Life Sci 53: 1597–1604, 1993.[CrossRef][ISI][Medline]
  15. Haller M, Brechtelsbauer H, Finsterer U, Forst H, Bein T, Briegel J, Peter K. The determination of plasma volume using indocyanine green in man. Anaesthesist 41: 115–120, 1992.[ISI][Medline]
  16. Haneda K, Horiuchi T. A method for measurement of total circulating blood volume using indocyanine green. J Exp Med 148: 49–56, 1986.
  17. Henry CB, Duling BR. Permeation of the luminal capillary glycocalyx is determined by hyaluronan. Am J Physiol Heart Circ Physiol 277: H508–H514, 1999.[Abstract/Free Full Text]
  18. Hunton DB, Bollman JL, Hoffman HN. Studies of hepatic function with indocyanine green. Gastroenterology 39: 713–724, 1960.[ISI][Medline]
  19. International Committee for Standardization in Haematology. Recommended methods for measurement of red-cell and plasma volume. J Nucl Med 21: 793–800, 1980.[Free Full Text]
  20. Ishihara H, Okawa H, Iwakawa T, Umegaki N, Tsubo T, Matsuki A. Does indocyanine green accurately measure plasma volume early after cardiac surgery? Anesth Analg 94: 781–786, 2002.[Abstract/Free Full Text]
  21. Jacob M, Bruegger D, Rehm M, Welsch U, Conzen P, Becker BF. Contrasting effects of colloid and crystalloid resuscitation fluids on cardiac vascular permeability. Anesthesiology 104: 1223–1231, 2006.[CrossRef][ISI][Medline]
  22. Jacob M, Rehm M, Orth V, Lotsch M, Brechtelsbauer H, Weninger E, Finsterer U. Exact measurement of the volume effect of 6% hydoxyethyl starch 130/0.4 (Voluven) during acute preoperative normovolemic hemodilution. Anaesthesist 52: 896–904, 2003.[CrossRef][ISI][Medline]
  23. Jones JG, Wardrop CA. Measurement of blood volume in surgical and intensive care practice. Br J Anaesth 84: 226–235, 2000.[Abstract/Free Full Text]
  24. Kisor DF, Frye RF, Kudsk KA. Estimation of the hepatic ratio of indocyanine green in swine. Clin Sci (Lond) 84: 681–685, 1993.[Medline]
  25. Kornbrust DJ, Barfknecht TR. Comparison of 7 azo dyes and their azo reduction products in the rat and hamster hepatozyte primary culture/DNA repair assays. Mutat Res 136: 255–266, 1984.[CrossRef][ISI][Medline]
  26. Massey EJ, de Souza P, Findlay G, Smithies M, Shah S, Spark P, Newcombe RG, Phillips C, Wardrop CA, Robinson GT. Clinically practical blood volume assessment with fluorescein-labeled HES. Transfusion 44: 151–157, 2004.[CrossRef][ISI][Medline]
  27. Masters AM, Leslie AJ, Kaldor I. Red cell and plasma volume development in newborn rats measured with double label. Am J Physiol 222: 49–54, 1972.[Free Full Text]
  28. Menth-Meier SE, Imoberdorf R, Regli B, Kipfer B, Turgay M, Ballmer PE. Determination of plasma volume by indocyanine green—validation of the method and use in patients after cardiopulmonary bypass. Intensive Care Med 27: 925–929, 2001.[CrossRef][ISI][Medline]
  29. Mi WD, Ishihara H, Sakai T, Matsuki A. Possible overestimation of indocyanine green-derived plasma volume early after induction of anesthesia with propofol/fentanyl. Anesth Analg 97: 1421–1427, 2003.[Abstract/Free Full Text]
  30. Najean Y, Deschrywer F. The body venous hematocrit ratio and its use for calculating total blood volume from fractional volumes. J Nucl Med 9: 558–560, 1984.
  31. Nakane H, Balzarini J, De Clercq E, Ono K. Differential inhibition of various deoxyribonucleic acid polymerases by Evans blue and aurintricarboxylic acid. Eur J Biochem 177: 91–96, 1988.[ISI][Medline]
  32. O'Brien JJ, Heath TJ. Effect of anaesthesia with cyclopropane, halothane or thiopentone on the vascular compartment in sheep. Br J Anaesth 40: 853–863, 1968.[Abstract/Free Full Text]
  33. Orth VH, Rehm M, Thiel M, Kreimeier U, Haller M, Brechtelsbauer H, Finsterer U. First clinical implications of perioperative red cell volume measurement with a nonradioactive marker (sodium fluorescein). Anesth Analg 87: 1234–1238, 1998.[Abstract/Free Full Text]
  34. Ott P, Weisinger RA. Nontraditional effects of protein binding and hematocrit on uptake of indocyanine green by perfused rat liver. Am J Physiol Gastrointest Liver Physiol 273: G227–G238, 1997.[Abstract/Free Full Text]
  35. Paumgartner G. The handling of indocyanine green by the liver. Schweiz Med Wochenschr 105 Suppl 17: 1–30, 1975.[ISI][Medline]
  36. Pearson TC, Guthrie DL, Simpson J, Chinn S, Barosi G, Ferrant A, Lewis SM, Najean Y. Interpretation of measured red cell mass and plasma volume in adults: Expert Panel on Radionuclides of the International Council for Standardization in Haematology. Br J Haematol 89: 748–756, 1995.[ISI][Medline]
  37. Pries AR, Secomb TW, Gaehtgens P. The endothelial surface layer. Pflügers Arch 440: 653–666, 2000.[CrossRef][ISI][Medline]
  38. Regillo CD. The present role of indocyanine green angiography in ophthalmology. Curr Opin Ophthalmol 10: 189–196, 1999.[CrossRef][Medline]
  39. Rehm M, Haller M, Brechtelsbauer H, Akbulut C, Finsterer U. Changes in plasma volume in immediate pre- and postoperative periods in patients with major gynecologic surgery. Infusionsther Transfusionsmed 25: 222–228, 1998.
  40. Rehm M, Haller M, Orth V, Kreimeier U, Jacob M, Dressel H, Mayer S, Brechtelsbauer H, Finsterer U. Changes in blood volume and hematocrit during acute preoperative volume loading with 5% albumin or 6% hetastarch solutions in patients before radical hysterectomy. Anesthesiology 95: 849–856, 2001.[CrossRef][ISI][Medline]
  41. Rehm M, Orth V, Kreimeier U, Thiel M, Maller M, Brechtelsbauer H, Finsterer U. Changes in intravascular volume during acute normovolemic hemodilution and intraoperative retransfusion in patients with radical hysterectomy. Anesthesiology 92: 657–664, 2000.[CrossRef][ISI][Medline]
  42. Rehm M, Orth V, Kreimeier U, Thiel M, Haller M, Brechtelsbauer H, Finsterer U. Four cases of radical hysterectomy with acute normovolemic hemodilution despite low preoperative hematocrit values. Anesth Analg 90: 852–855, 2000.[Free Full Text]
  43. Rehm M, Zahler S, Lotsch M, Welsch U, Conzen P, Jacob M, Becker BF. Endothelial glycocalyx as an additional barrier determining extravasation of 6% hydroxyethyl starch or 5% albumin solutions in the coronary vascular bed. Anesthesiology 100: 1211–1223, 2004.[CrossRef][ISI][Medline]
  44. Retzlaff JA, Tauxe WN, Kiely JM, Stroebel CF. Erythrocyte volume, plasma volume, and lean body mass in adult men and women. Blood 33: 649–661, 1969.[Abstract/Free Full Text]
  45. Rowland M, Benet LZ, Graham GG. Clearance concepts in pharmacokinetics. J Pharmacokinet Biopharm 1: 123–136, 1973.[CrossRef][ISI][Medline]
  46. Sawka MN, Young AJ, Pandolf KB, Dennis RC, Valeri CR. Erythrocyte, plasma, and blood volume of healthy young men. Med Sci Sports Exerc 24: 447–453, 1992.
  47. Schad H, Haider M, Brechtelsbauer H. Determination of plasma volume with indocyanine green. Anaesthesist 36: 608–614, 1987.[ISI][Medline]
  48. Sekimoto M, Fukui M, Fujita K. Plasma volume estimation using indocyanine green with biexponential regression analysis of the decay curves. Anaesthesia 52: 1166–1172, 1997.[CrossRef][ISI][Medline]
  49. Speich R, Saesseli B, Hoffmann U, Neftel KA, Reichen J. Anaphylactoid reactions after indocyanine green administration. Ann Intern Med 109: 345–346, 1988.[ISI][Medline]
  50. Thomas E, Jones G, de Souza P, Wardrop C, Wusteman F. Measuring blood volume with fluorescent-labeled hydroxyethyl starch. Crit Care Med 28: 627–631, 2000.[CrossRef][ISI][Medline]
  51. Tschaikowsky K, Meisner M, Durst R, Rugheimer E. Blood volume determination using hydroxyethyl starch: a rapid and simple intravenous injection method. Crit Care Med 25: 599–606, 1997.[CrossRef][ISI][Medline]
  52. Vande Walle J, Donckerwolcke R, Boer P, Van Isselt HW, Koomans HA, Joles JA. Blood volume, colloid osmotic pressure, and f-cell ratio in children with nephrotic syndrome. Kidney Int 49: 1471–1477, 1996.[ISI][Medline]
  53. Vink H, Duling BR. Capillary endothelial surface layer selectively reduces plasma solute distribution volume. Am J Physiol Heart Circ Physiol 278: H285–H289, 2000.[Abstract/Free Full Text]
  54. Vink H, Duling BR. Identification of distinct luminal domains for macromolecules, erythrocytes, and leukocytes within mammalian capillaries. Circ Res 79: 581–589, 1996.[Abstract/Free Full Text]
  55. Vogel J, Sperandio M, Pries AR, Linderkamp O, Gaehtgens P, Kuschinsky W. Influence of the endothelial glycocalyx on cerebral blood flow in mice. J Cereb Blood Flow Metab 20: 1571–1578, 2000.[CrossRef][ISI][Medline]



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M. Rehm, D. Bruegger, F. Christ, P. Conzen, M. Thiel, M. Jacob, D. Chappell, M. Stoeckelhuber, U. Welsch, B. Reichart, et al.
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