Stability of perfluorocarbon blood substitutes determined by sedimentation field-flow fractionation

ANALYTICAL

BIOCHEMISTRY

138,488-494

(1984)

Stability of Perfluorocarbon Blood Substitutes Determined by Sedimentation Field-Flow Fractionation FENG-SHYANGYANG,KARIND.CALDWELL,J.CALVINGIDDINGS,ANDLYNNASTLE* Department of Chemistry and *Department of Medical Technology, College of Pharmacy, University of Utah, Salt Lake City, Utah 841 I2 Received October 14, 1983 It is shown that the method known as sedimentation field-flow fractionation, which has been applied to the separation and characterization of many industrial and biological particles and recently to emulsions, can be used to obtain high-resolution dropiet diameter profiles for perfluorocarbon blood substitutes. Following a description of the methodology, experiments are described for two commercial pertluorocarbon emulsions, Fluosol-DA 20% and Fluosol-43. The droplet diameter profiles for both of these blood substitutes are shown to shift to noticeably higher diameter values in less than 2 months. The diameter at the profile peak for Ruosol-DA 20’S, for example, shifts from 0.19 to 0.27 pm in 56 days. KEY WORDS: emulsion stability; blood substitutes; artificial blood; field-flow fractionation; emulsion sizing; particle characterization.

Emulsions of pet-fluorocarbons have received considerable attention as gas transporting agents in blood substitutes because of the high solubility of gases (approximately 3050 ml Oz/ml and 150-230 ml COz/ml at 760 torr) in per-fluorocarbons. Such emulsions not only can serve as nonantigenic resuscitation fluids, but the small size (~0.2 pm) of the emulsion droplets, decreased viscosity and decreased surface tension due to surfactants, have also stimulated research into their effectiveness in transporting oxygen past cerebral (l-3) and myocardial (4,5) emboli. Perfluorocarbon emulsions have been investigated extensively as perfusion fluids for organ preservation (6-8) and they also hold promise for reducing morbidity of hemoglobin-binding substances such as carbon monoxide and cyanide (9,10) and in sickle cell crises. Historically, the success of perhuorocarbon blood replacement and exchange transfusions has been dependent upon the preparation of fine, stable emulsions. Geyer and co-workers (11,12) were the first to totally replace the blood of rats with an emulsion of perfhtoro-

ooo3-2697/84 $3.00 Copyright 0 1984 by Academic F’rcs, Inc. All rigbts of nproduclion in any form -cd.

tributylamine (FC-43)’ and maintain them for short periods of time in an oxygen atmosphere. With more finely dispersed and stable emulsions animals survived and could be removed from the oxygen atmosphere .after the third day (13), and with further improvements in the emulsion (14) and the fluid medium ( 1516) survival and morbidity were improved. The persistence of an emulsion in the circulation is dependent upon the size of the emulsion particles, the nature of the perfluorochemical, and the surface characteristics. Not only are finely dispersed emulsions more stable than coarse dispersions, but larger particles are clear& from the circulation more rapidly than smaller ones (17). Thus, the most important characteristic of a perfluorocarbon blood substitute is the particle size and stability oftheemulsionintbeb~(14,16,18). A concern of secondary importance is elimination of the pet-fluorocarbon. The reticuloendothelial system will remove the emulsion ’ Abbreviations used: FC-43, pertluorotributylamine; FFF, field-flow fractionation.

488

STABILITY

OF BLOOD

SUBSTITUTES

489

dropletsfrom thecirculation,but theprincipal Unfortunately, there Tao methods in the routeof elimination from the body is through standard stable of partisre-sizing techniques the lungs (14). The log of the rate of elimination is directly related to the vapor pressure and inversely related to the boiling point ( 14). FC-43 forms very stable emulsions which function well in blood substitutes, but the perfluorocarbon persists indefinitely in the spleen and liver and is found in vacuoles lilled with the perlluorocarbon in hepatocytes, Kupffer cells, and histiocytes (19-2 1). Another compound which has been utilized extensively is pertluorodecalin, which has a vapor pressure low enough not to cause gas embolism, yet high enough to be eliminated through the skin and lungs in a few days (14,19,22). However, its emulsions are not consistent and have performed poorly. A large number of fluorocarbons have been evaluated for possible use as blood substitutes (23,24). The Green Cross Corporation of Japan has made two perfluorocarbon emulsion preparations commercially available for research. The first is Fluosol-43, an emulsion of FC-43 in a balanced salt solution with hydroxyethyl starch for oncotic activity and Pluronic F-68, a polyoxyethylene surfactant. The second is Fluosol-DA, a 20% (w/v) emulsion of perfluorodecalin in a similar medium with perfluorotripropylamine and egg yolk phospholipids added to stabilize the emulsion. FluosolDA has been administered to normal human volunteers (25) and has been used in therapeutic clinical studies in larger numbers of patients ( 10,26-28) without significant adverse effects (29,30). However, even with these encouraging results the problem of emulsion stability during storage and in vim still persists. Considerable effort is still being expended to develop more stable emulsions from perlluorocarbons which will be readily excreted. Because the emulsion stability is the most important factor in the development of a suitable blood substitute, it would be advantageous to have a method to monitor the droplet size distribution so that this problem can be studied and dealt with.

which work effectively for these emulsions. Electron microscopy is applicable to emulsions only with heavy-metal staining, which is not always easily performed (3 1). Couher counting methods and light microscopy do not work effectively below 0.5~pm diameter. Liit-scattering methods do not provide the significant details of the droplet diameter distribution. A relatively new technique, sedimentation field-flow fractionation (sedimentation FFF), has the capability of separating particles according to size (32,33). The method has been shown to provide detailed size distribution data for many kinds of industrial and biological particles (34,35). Recently, sedimentation FFF was shown to be applicable to emulsions (36). This work would suggest that changes in the size distribution of perfluorocarbon emulsions could be followed accurately and in detail by using sedimentation FFF. METHODOLOGY

Field-flow fractionation is a high-resolution separation technique which resembles chromatography with respect to its experimental sequence (37). A narrow band of sample is injected into a flowing carrier stream via a sample injection system, carried through a column where fractionation takes place, and then led into a detector and possibly a fraction collector, if collected fractions are desired. The main difference between FFF and chromatography lies in the nature of the sep aration column. In FFF, the column consists of a narrow ribbon-shaped flow channel across which an external field is applied. There is no adsorptive or partitioning medium in the channel, which is entirely open and unobstructed. Rather than using a retentive medium, separation is induced by an external field or gradient applied perpendicular to the channel walls and thus to the flow direction. The technique is therefore also unlike electrophoresis and centrifugation techniques in

490

YANG

the sense that the field is applied in a direction perpendicular rather than parallel to the direction along which separation is occurring. The external field forces sample particles toward one wall, where, because of the viscous drag of the wall, the carrier stream which entrains the particles is traveling at a less than average velocity. As different particles react differently to the field, they will form more or less compact layers near the wall. In sedimentation FFF the applied gravitational field exerts a force on an injected sample which is proportional to particle mass. Big particles, which are forced into closer proximity with the wall than small particles, are therefore carried downstream more slowly than small particles. This is the origin of the differential migration of the sample and the consequent highresolution separation of its different-sized components. The rate of migration can be described by rigorous theoretical expressions, which allow one to convert the response of the detector to the sequence of emerging particles into a particle size profile. The theory of FFF has been presented in many places and will not be repeated here (32,33). An essential conclusion of the theory is that the retention volume V, or retention time tr of a homogeneous population of particles can be related to a basic retention parameter X

vr -=-= v”

tr to

1 6X(coth (1/2X) - 2X) ’

[II

in which X depends on the kind of field applied, the strength of that field, and particle characteristics such as density and diameter. In the present case, where the channel system is wrapped inside a centrifuge basket, the applied sedimentation field forces particles to move radially and accumulate at one of the channel walls. If the particle is denser than the channel fluid, the accumulation occurs at the outer wall of the channel, whereas a less dense sample moves toward the inner wall. The retention equations remain the same regardless of the accumulation wall due to the

ET

AL.

total symmetry of the system. The accumulation process is opposed by diffusion, and at equilibrium there is a complete balance between these two fluxes. The thickness of the resulting equilibrium distribution is given by the dimensionless parameter X, which for sedimentation FFF is related to the particle or droplet diameter d and the strength of the sedimentation field G (expressed in units of acceleration) by

VI where k is Boltzmann’s constant, T is temperature, w is channel thickness, and Ap is the absolute value of the difference between the particle density and the carrier density. Sedimentation field strength G can be expressed as G = ro(2nrpm/60)2, [31 where r. is the distance from the axis of centrifugation to the flow channel and “rpm” is the spin rate in revolutions per minute. Equation [2] shows that X varies with d, and Eq. [ 1] shows that V, and tr depend on X. Therefore, retention volume and time depend in a calculable way on droplet diameter d, such that a fractogram representing the emerging sequence of increasing droplet sizes can be converted into a detailed droplet diameter profile. EXPERIMENTAL

Emulsions. The two fluorocarbon emulsions studied, Fluosol-43 and Fluosol-DA 20%, were gifts from Alpha-Therapeutics Company of Los Angeles, California. These emulsions are products of the Green Cross Corporation, Joto-ku, Osaka, Japan. The densities of perfluorotripropylamine and perfluorodecalin are 1.80 and 1.94 g/cm3, respectively (Dr. Charles Heldebrant, AlphaTherapeutics, private communication). For evaluation of the droplet size distribution we assume the average density of the droplets in the emulsions to be 1.9 g/cm3. The main com-

STABILITY

OF BLOOD

SUBSTITUTES

491

ponent of the Fluosol-43 emulsion droplets is (38,39). Following ,this compromise, a field perlluorotributylamine (20% w/v), while the strength of 86.2g was found suitable for FluoFluosol-DA 20% emulsion droplets are comsol-43 and 21.5g for Fluosol-DA 20%. The posed of a mixture of perfluorodecalin (70% flow rate used in both cases was 60 ml/h. w/v) and perfluorotripropylamine (30% w/v). Pluronic F-68 (~2.6%) served as emulsifier RESULTS AND DISCUSSION in both cases, assisted by egg yolk phospholipids in the case of Fluosol-DA 20% (0.4%). The top curve of Fig. 1 is a fractogram of The fluorocarbons were emulsified in balanced Fluosol-DA 20% recorded on the day the salt solutions with hydroxyethyl starch for on- emulsion was prepared. The figure shows that cotic activity suitable for intravenous injec- the maximum of the emulsion distribution tion. emerged within 10 min of the start of flow. The fractograms are highly repeatable from To initiate the stability tests, the fluorocarbon emulsions were prepared according to the run to run. instructions provided by the manufacturer. The use of Eqs. [I] and [2] makes possible During the test period, the emulsions were the calculation of the diameter d of the narrow stored at 4°C. population of droplets that are eluted at any Apparatus. The general experimental pro- specific retention volume V, or retention time cedure and apparatus for FFF have been de- tr. Therefore, these equations can be used to scribed previously (32). In the present exper- construct a droplet diameter scale along the iments the FFF channel dimensions were 79.4 axis of Fig. 1. This scale is shown at the top X 2.0 X 0.0127 cm3, giving a channel void of the figure. It is seen that the size distribution volume of 2.0 ml (determined by elution of of freshly prepared (Day 0) Fluosol-DA 20% a nonretained sample). The channel was reaches a maximum just below the droplet curved to fit inside a centrifuge rotor basket diameter of 0.2 pm and then falls rapidly, in which the ribbon-like channel was positioned 7.7 cm from the axis of rotation. Droplet diheter tp-4 Procedure. The carrier liquid (phosphate al 0.2 0.3 0.35 0.4 buffer solution, 0.1 M, pH 7) was fed into the I FFF system at a constant flow rate by means of a Gilson Minipuls 2 peristaltic pump. The channel effluent was monitored for absorbance at 253.7 nm by an LDC Model 1205 uv detector with a cell volume of 8 ~1. The detector signal was fed to a lo-in. flat-bed Omniscribe recorder from Houston Instruments. Ten microliters of each emulsion sample was injected by syringe at the head of the FFF Retentkm time (mid channel. Before each sampling, the container was shaken for 10 s. FIG. 1. Time variation of fmctogmms for Fluosol-DA After injection the channel flow was stopped 20% by sedimentation FFF. The ordinate measures the and the spin started. The sample was allowed detector response at 253.7 pm. (The major part of this to relax to its equilibrium distribution for 10 response is due to light scattering, which is particle size min under the influence of the chosen cen- dependent. Thus, the trace represents a slightly distorted size distribution. A second distortion arises from trifugal field before the flow was resumed. The droplet the different resolution of different particle diametersfield strength (spin rate) was selected low see text.) Field = 21.Q flow = 60 ml/h; V” = 2.0 ml; enough to avoid significant steric effects sample = 10 pl of Fluosol-DA 20%.

YANG

492

ET AL. Dr@tdmmeler

TABLE I SIZE VARIATION OF DROPLETS OF FLUO~~L-DA AT PEAK MAXIMUM WITH ELAPSED TIME Day

WV,

d (crm)

0

0.250 0.247 0.221 0.181 0.201 0.181 0.163 0.168 0.193 0.165 0.159 0.100 0.092

0.192 0.192

I 2 4 I

9 13 17 19 28 41 51 56

20%

0.201 0.216 0.208 0.216 0.224 0.222 0.211 0.223 0.226 0.266 0.274

declining to only 10-l 5% of its peak value at 0.3 pm. This result confirms that the droplet diameter distributions of the freshly prepared emulsion are quite narrow. With aging, there is a clearly discernible trend toward larger particle sizes in the size distribution of the Fluosol-DA 20%. The changes are followed in some detail in Table 1, where the results of fractograms acquired on 13 different days-spanning the range from Day 0 to Day 56-are summarized. The final (Day 56) fractogram, presented as the bottom curve of Fig. 1, shows how extreme the changes are in that interval; the droplet diameter at the peak has increased by over 40%, and the shift to a wider distribution was also observed, as is apparent from the figure. The gradual and subtle shifts indicated by the fractograms reflect themselves visually after Day 41, at which time a noticeable sediment was observed in the sample. Fluosol-43 is composed of smaller droplets than Fluosol-DA 20%, as shown by the top fractogram of Fig. 2. This fmctogram shows that the droplet diameter of the freshly prepared emulsion at the peak maximum is about 0.12 pm. However, a shift in the droplet size distribution with time to larger droplets is ap-

I

-e----

0.1 0.15 111’~1’1~11~

(,um)

0.2

3 1

30 Retention time hinl

’

0.25 I

r

60

FIG. 2. Time variation of fractograms for Flusol- by sedimentation FFF. The ordinate measures detector response at 253.7 km. Field = 86.2~ flow = 60 ml/h; P = 2.0 ml; sample = 10 ~1 of Fhwsol-43.

parent for Fluosol-43, as for the companion emulsion. This shift is detailed in Fig. 2 and Table 2. The change, although followed only to Day 36, does not appear to be so rapid as that for Fluosol-DA 20%. The validity of the droplet diameter pro&es is difficult to judge in absolute terms because no other methods have been found to provide this information. However, self-consistency tests, in which essential FFF conditions are changed and the results compared, can provide strong support for the method. Extensive preTABLE 2 SUE VARIATIONS OF DROPLETS OF F~uosoL-43 PEAK MAXIMUM WITH ELAPSED TIME

DaY

R = V”/V,

d (4

0 1 4 8 12 14

0.235 0.234 0.229 0.224 0.197 0.200 0.215 0.188 0.210 0.199

0.124 0.124 0.125 0.126 0.132 0.132 0.128 0.134 0.129 0.132

19 23 29 36

AT

STABILITY

OF BLOOD

vious tests with emulsions, employing changes in spin rate, flow velocity, sample size, and the FFF column itself, have shown excellent self-consistency (36). We regard changes in spin rate (field strength) as providing the most rigorous of comparisons, and have employed such changes with the present emulsions. If there are any significant errors in our results caused by droplet-wall interactions, dropletdroplet interactions, including aggregation, tinite droplet diameter (steric) effects, or intrinsic band broadening in the FFF column, there should be a significant divergence of results obtained at two different spin rates. Our experiments show a difference of less than 6% in the diameter at the size profile maximum for Fluosol-DA 20% when compared at field strengths of 2 1.5 and 42.2g. A difference of less than 8% applied to Fluosol43 when run at field strengths of 42.2 and 86.2g. The difference near the profile maximum was larger than that elsewhere in the size profile in both cases. The integrity of both emulsions was therefore believed to be well maintained under the applied field strengths used in the stability studies (2 1Sg for FluosolDA 20% and 86.2g for Fluosol-43), and the overall results confirmed the basic correctness of the size profiles. We note that the curves we have shown in Figs. 1 and 2 are actually “fractograms,” which show a detector’s response to the sired droplet sequence emerging from the FFF+column. Inasmuch as the particles are sorted according to size and our theory permits us to construct a droplet diameter scale along the time axis, the fractogram can be referred to as a droplet diameter profile, an expression used above. While these profiles reflect size changes, and can therefore accurately trace aging effects or manufacturing differences, they are not quantitative droplet size distribution curves as such. The fractogram must be corrected by a scale factor and a detector response factor in order to arrive at a true droplet size distribution curve. While the techniques for these corrections have been previously applied to emul-

493

SUBSTITUTES

sions (36) and could be extended to the present fluorocarbon emulsions if certain optical properties of the emulsions were determined, the corrections are deemed unnecessary for the purpose of following the changes in size characteristics with time or manufacturing process. Finally, we have not discussed the physicochemical reasons for the droplet size increase observed with aging. This could involve either aggregation or the slow growth of large droplets at the expense of small droplets by virtue of slight solubility differences in the aqueous phase. It is believed that the effects of the two mechanisms could be sorted out by comparing the predictions of theoretical models with the detailed shifts in actual size characteristics made available by FFF. Such studies should eventually lead to means for improving emulsion stability and for monitoring emulsion stability and persistence in vivo. ACKNOWLEDGMENT This investigation was supportedby Public Health Service Grant GM 1085 1-25 from the National Institutes of Health.

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