Iowa State University Capstones, Theses and Retrospective Theses and Dissertations Dissertations
1981 Evaluation of p-aminohippurate as a marker for blood-flow determinations in cattle Denis Jay Meerdink Iowa State University
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University Microfilms International 300 N. ZEEB RD., ANN ARBOR, Ml 48106 8209148
Meerdink, Denis Jay
EVALUATION OF P-AMINOHIPPURATE AS A MARKER FOR BLOOD- FLOW DETERMINATIONS IN CATTLE
Iowa State University PH.D. 1981
University Microfilms I nt6r n âtion&! 300 N.Zeeb Road, Ann Arbor, MI 48106 Evaluation of p-aminohippurate as a marker
for blood-flow determinations in cattle
by
Denis Jay Meerdink
A Dissertation Submitted to the
Graduate Faculty in Partial Fulfillment of the
Requirements for the Degree of
DOCTOR OF PHILOSOPHY
Department: Animal Science Major: Nutritional Physiology
Approved: Members of the Committee:
Signature was redacted for privacy. Signature was redacted for privacy.
Signature was redacted for privacy.
Signature was redacted for privacy.
Iowa State University Ames, Iowa
1981 11
TABLE OF CONTENTS
Page
PREFACE 1
Format of Dissertation 2
PART I. DETERMINATION OF PARA-AMINOHIPPURIC ACID AND N-ACETYL-PARA-AMINOHIPPURIC ACID BY HIGH PRESSURE LIQUID CHROMATOGRAPHY 4
Introduction 4
Literature Review 4
Materials and Methods 7
Materials 7 Methods 8 Detection wavelength 8 Chromatographic conditions 10 Detector linearity 10 Deprotelnlzatlon 11 Barium hydroxide-zinc sulfate 11 Perchloric acid-potassium hydroxide 12 Peak-height evaluation 14
Results and Discussion 15 Detection wavelength 15 Chromatographic condlt^on^ 15 Detector linearity 18 Deprotelnlzatlon 18 Barium hydroxide-zinc sulfate 21 Perchloric acid-potassium hydroxide 22
Conclusions 27
PART II. PARA-AMINOHIPPURIC ACID IN VIVO 29
Literature Review 29
Some techniques for the determination of flow 29 Electromagnetic flowmeters 29 Ultrasound flowmeters 30 Thermal methods 31 Other methods 32 General limitations 34 Determination of blood flow by indicator-dilution 34 ill
Page
Renal transport of PAH 37 PAH clearances and kinetics in vivo 39 Metabolism of PAH 43
Materials and Methods 45
Experiments with non-radioactive PAH and aPAH 45 Single injections 45 Continuous infusions 48 Experiments with radioactive PAH 49
Results and Discussion 53
Experiments with non-radioactive PAH and aPAH 53 Single injection 53 Continuous infusion 55 General comments on the non-radioactive experiments. 56 Experiments with H-PAH 60
Conclusions 85
POSTSCRIPT 87
BIBLIOGRAPHY 88
ACKNOWLEDGEMENTS 98
APPENDIX A. PEAK-HEIGHT RATIO PROCEDURE FOR QUANTITATION OF PAH AND aPAH USING PAB AS AN INTERNAL STANDARD 99
APPENDIX B. EVANS BLUE PROCEDURE 101
APPENDIX C. PLOTS FOR SINGLE-INJECTION EXPERIMENTS USING UNLABELLED PAH AND aPAH 103
APPENDIX D. PLOTS FOR CONTINUOUS INFUSION EXPERIMENTS USING UNLABELLED PAH AND aPAH 158 1
PREFACE
In recent years, much knowledge has been obtained about metabolism by investigating the in vivo kinetics of radioactively-labeled compounds.
Much information is lacking, however, about changes of various compounds in metabolic diseases such as ketosis and about the role that specific organs have in altered metabolic states occuring in such diseases.
Although concentrations of metabolites can be measured in blood and information is available regarding in vitro metabolic capacities, the physiological changes in metabolism cannot be described with confidence until information is obtained under physiological conditions. One way to assess the changes of metabolic adaptation is by trans-organ balances of
Important metabolites. The trans-organ balance approach estimates fluxes of various metabolites in and out of organs of Interest by comparing blood entering and exiting the organ. The trans-organ balance method reveals changes in organ uptake and output of a metabolite regardless of a change, or lack of change, in systemic blood concentrations of that metabolite.
A requirement of the trans-organ balance technique is an accurate quantitation of flow of blood that enters and exits organs to be studied.
Fluxes are calculated from concentrations of the metabolite in blood and the rate of flow of blood through the organ. Therefore, small errors in determination of blood flow introduce errors in estimates of fluxes in spite of an accurate determination of metabolite concentration in blood.
Dye-dilution techniques have been used successfully to determine flow of blood in vivo. These techniques involve Injection of an indicator or 2
marker and determination of dilution of that marker in sampled blood.
Continuous-infusion techniques have an advantage for use in trans-organ
balance studies in that the timing of collection of samples is not critical.
Therefore, substantial errors in calculation of fluxes across organs can
be avoided.
An acceptable blood-flow marker must exhibit four qualities:
(1) be rapidly and uniformly distributed in blood, (2) have no
physiological or pharmacological action on the cardiovascular system
that would affect the flow of blood, (3) be neither produced nor de
stroyed within the blood stream, and (A) be easily detected.
It was of interest to the Nutritional Physiology Group in the
Department of Animal Science at Iowa State University to develop a dye-
dilution method for determination of blood flow in experiments investi
gating hepatic fluxes of metabolites in ruminant metabolism including
ketosis, a metabolic disease common to lactating dairy cows.
Format of Dissertation
The research described in this dissertation has been divided into two
parts. This two-part format is an attempt to present and discuss two distinct segments or goals of the research undertaken to evaluate para- am inohippurate (PAH) as a blood-flow marker to be used for trans-organ
balance studies. 3
Part I is concerned with the development of an accurate, rapid method for quantitation of PAH and N-acetyl-p-aminohippurate (aPAH).
Part II involves the in vivo observations of the kinetics and trans formations of PAH and aPAH. Each part contains its own Literature Review,
Materials and Methods, Results and Disccusion, and Conclusion sections. 4
PART I.
DETERMINATION OF PARA-AMINOHIPPURIC ACID AND
N-ACETYL-PARA-AMINOHIPPURIC ACID BY
HIGH PRESSURE LIQUID CHROMATOGRAPHY
Introduction
Para-amlnohippuric acid is well-suited for use as a blood-flow marker in trans-organ balance studies, especially for trans-liver balances, because it is not extracted by the liver (Katz & Bergman, 1969a) and is excreted only by the kidney (Blaufox, et al., 1963). Problems associated with analysis available for PAH needed to be overcome by development of a new analysis procedure. The problems will become apparent in the litera ture review that follows.
Literature Review 4 PAH is an N -amino-substituted derivative of hippuric acid, and has been used for a variety of purposes in renal physiology and clinical medicine. The parent compound was termed "hippuric acid" after the site of its original discovery, horse urine.
PAH perhaps is most closely identified with renal physiology because of its traditional use in the determination of renal plasma flow and of
renal clearance. The first studies on excretion of hippuric acid deriv
atives were carried out durinp the late 1930s and early 1940s (Elsom, et al., 1936; Smith, et al., 1938; Finkelstein, et al., 1941; and Smith, et al., 1945). Inconsistencies in clearance values of diodrast
(3,4-diiodo-4-pyridone-N-acetate), another compound commonly used to 5
determine renal plasma flow and clearances, and of PAH prompted Newman, et al. (1949) to investigate some factors that might cause variation in clearances. Their study indicated that constant plasma clearances occur with decreasing plasma concentrations after single injections of PAH in dogs, as well as with N-acetyl-para-aminohippurate (aPAH) in dogs and man. Acetylated-PAH is the major metabolite of PAH in mammals; the conversion of PAH to aPAH will be discussed in Part II.
The clinical use of PAH has led to the developement of a number
of Methods for determining PAH in blood. Bratton and Marshall (1939)
developed a method for determination of sulfanilamide that Involved
coupling N-(l-napthyl) ethylenedlamine dihydrochloride to the para-amino
group, which formed an azo dye that could be determined colorimetrically.
Smith, et al. (1945) developed a widely used method by altering reagent concentrations of the Bratton and Marshall procedure. Newman, et al.
(1949) further adapted the method for use in studies on renal clearances.
Significant disadvantages exist with these methods, however, in that there
is a necessity for careful timing of adding reagents, a lack of reagent
stability, and a lack in the color permanence. More importantly, the methods lack specificity for PAH because many primary amines react
(Waugh & Beall, 1974).
Brun (1951) developed an assay that utilized formation of a Schiff
base upon reaction of p-dimethylaminobenzaldehyde with the primary
p-amino group of PAH. This assay requires deproteinization only in
samples that contain low concentrations of PAH and different sample 6
preparation for samples containing high and low concentrations.
Automated methods for analysis of PAH have been developed for
clinical use (Harvey & Brothers, 1962; Parekh, et al., 1974; and Looye,
1970). These methods are based on the coupling reactions described
already and potentially can give positive results for a large number of
primary amines. Selectivity of reagents has been the major problem with
all colorimetric analyses of PAH (Shoup & Kissinger, 1975).
With the advent and advances of high pressure liquid chromatography
(HPLC) in recent years, procedures have been developed to determine PAH
without coupling reactions in an attempt to avoid problems encountered
with colorimetric analyses. Shoup and Kissinger (1975) developed a
method that utilizes thin-layer electrochemical oxidation for detection
of PAH. Their method requires a specialized detection system that they
developed. In a method developed by Brown, et al. (1974), and later
Improved by Brown and Michalski (1976), PAH and aPAH were separated in
approximately 20 minutes and detected by UV absorbance at 254 nm. A
critical requirement was that the 5 mM formate buffer be pH 3.5. An
important advantage of their method, which will be discussed later, was
that PAH and aPAH were successfully separated and detected without
coupling reactions or pre-analysis hydrolysis.
Sensitivity is even more of a problem for detection of aPAH than
for PAH. With the colorimetric methods described already, the coupling
reagent reacts with primary aromatic amines to form either an azo dye 7
(the Bratton-Marshall method and subsequent modifications) or a Schiff base (Brun, 1951). However, the p-amino group of aPAH is a secondary amine and is therefore unreactive to coupling reagents. Bratton and
Marshall (1939) de-acetylated aPAH, to form PAH, by acid hydrolysis with
10 N hydrochloric acid at approximately 96°C for 3.5 hours. Brown, et al.
(1976) investigated the Bratton-Marshall hydrolysis procedure with HPLC, and found that conversion of aPAH to its de-acetylated (PAH) and de- glycinated (N-acetyl-para-aminobenzoic acid) forms is directly related to hydrolysis reaction time and concentration of hydrochloric acid. If either time or acid concentration were varied, differing amounts of arti facts were produced, which were not detectable by standard colorimetric procedures. They concluded that when colorimetric, chemical, or separation techniques are to be employed after hydrolysis, consideration must be given to secondary decomposition products that may or may not be reactive to coupling agents.
It is obvious that a method of analysis was needed that did not require acid hydrolysis for detection of aPAH so that questionable pre-analysis treatment could be avoided. Also, there was a need for development of a procedure that would quantitate PAH and aPAH directly to avoid non-specificity of colorimetric procedures.
Materials and Methods
Materials '
A Waters Associates (Milford, Mass.) HPLC system equipped with a 8
model 6000A pump, a U6K continuous-flow injector, and an RCM-100 radial 13 compression module containing a MicroBONDAPAK-C Radial-PAK cartridge was used for separations. A Perkin-Elmer (Oak Park, 111.) model 55 variable-wavelength spectrophotometer with a 20 microliter flow-through cuvette was used for detection. Recordings were made with a Houston
Instruments (Austin, Texas) Omniscribe recorder set at 0.1, 1, or 10 mV, respectively, providing 0.003, 0.03 or 0.3 absorbance units full scale.
Lyophilized PAH was obtained from Sigma Chemical Co. as the free acid.
Purchase of p-(glycyl-l-^** C)-aminohippuric acid was from New England
Nuclear (Boston, Mass.). N-acetyl-p-aminohippuric acid and N-acetyl-p-
(glycyl-l-^''C)-aminohippuric acid was prepared from PAH and ^C-PAH, respectively (Smith, et al., 1945). Purity was monitored by HPLC and by a melting point between 198 and 200°C. Para-aminobenzoic acid (PAB) was obtained from J. T. Baker Chemical Co.
Methods
Detection wavelength The choice of detection wavelength was based on spectral analysis of PAH, aPAH, and PAB. PAB was included so that its use as an internal standard could be evaluated, and was chosen because of its similarity in molecular structure to PAH and aPAH, thus suggesting a similarity in chromatographic behavior and absorbance. The molecular structures of PAH, aPAH, and PAB are in Figure 1.
Aqueous solutions of 1 mM PAH, aPAH, and PAB were prepared by dissolving 194.2 rag of PAH, 237.2 mg aPAH, and 137.1 mg of PAB in approxi- a M P V P Os C~N~CH2~C\ C-N-CHgC^ C-OH OH OH
O
NH, I^H NH c=o 2 CHc
PAH aPAH PAB
Figure 1. Molecular structures of PAH, , and PAB 10
mately 950 ml of de-ionized distilled water in separate beakers. The pH of each solution was adjusted to 3.5 with concentrated phosphoric acid, and
final dilution made to 1000 ml with additional water. The solutions then
were scanned separately from 200 to 300 nm for absorbance in quartz
cuvettes with a Beckman scanning spectrophotometer. The plots of
absorbance versus wavelength were superimposed and are presented in Results
and Discussion that follows.
Chromatographic conditions A reverse-phase chromatography system
was used to exploit differences in polarity of the three compounds in an
acidic medium. The conditions reported by Bush, et al. (1979) for
separation of volatile fatty acids were used as a starting point. The concentrations of buffer and methanol in the mobile phase were altered until a separation with baseline resolution of PAH, aPAH, and PAB was achieved in approximately 3 minutes. The mobile phase of 30% methanol
(V/V) in 10 mM NaH.PQ. buffer was mixed and the pH adjusted to 3.5 with 2. 4 concentrated phosphoric acid. Degassing and particle removal was by vacuum filtration through a 0.45 ym Millipore filter (FH type, moistened
with methanol). A flow rate of 2.5 ml/minute produced column pressures of 800 to 1000 psi. Standard solutions of PAH, aPAH, and PAB were pre
pared and adjusted to pH 7 or greater with NaOH before injection on the
HPLC. Absorbances of the three compounds were monitored at 270 nm and were recorded as absorbance versus time.
Detector linearity Linearity of detector response was evaluated by injecting three replicate 25 yl aliquots of two independently-prepared standard solutions containing varied concentrations of PAH, aPAH, or PAB. 11
The pH of these solutions was adjusted to approximately 7 with NaOH prior
to injection. Peak heights were measured from baseline and averaged
within each concentration for both sets of standard solutions. Average
peak heights were plotted versus nanomoles injected for each compound and
the least-squares estimate of linear equations calculated using the
Statistical Analysis System (SAS Institute, Inc., 1979).
Deproteinization Deproteinization agents for whole blood were
evaluated by recovery experiments using (glycyl-l-^^C)-PAH and (glycyl-1- 14 C)-aPAH. Triplicate solutions were prepared according to the following
summaries and the radioactivities compared among the treatments.
Barium hydroxide-zinc sulfate Ba(0H)2 and ZnSO^ were used
according to the procedure reported for blood glucose by Somogyi (1945). 14 The C-PAH standard was an aqueous solution containing 100 yg of PAH per
ml and having an activity of approximately 1 yCi/ml. The treatments were
prepared as outlined in Table 1.
Table 1. Treatments for evaluation of BaXOH^g-ZnSO^
1 M Water Blood Ba(OH)- Treatment (ml) (ml) (mi) ^
Control/Control 4 _ _
Control (C) 2-2
Blood -22 (B)
â l4f All treatments contained 60 yl of the C-PAH standard solution and were prepared in triplicate. 12
The Control and Blood treatments were mixed well and 2 ml of ZnSO^
slowly added during vortex stirring. The Control/Control treatment was
mixed well and 2 additional ml of water were added. The ZnSO^ solution
was balanced with the BaCOH)^ solution; i.e., when equal volumes of
isomolar Ba(0H)2 and ZnSO^ solutions were added, the resulting mixture
had a pH of approximately 7.
All three treatments were centrifuged for 20 minutes at approximately
13,000 X gravity and supernatants filtered through tissue paper. Aliquots
of 25 yl were injected onto the HPLC system and the eluent collected during
the 270 nm absorbance peak for PAH. To the collected eluents, 15 ml of
Ready-Solv (EP type from Beckman) were added, and the samples were counted on a Beckman 8000 liquid scintillation spectrophotometer. Observed CPM were converted to DPM by using the counting efficiency calculated from
"H" numbers. DPM were averaged within each treatment and summarized in the following Results and Discussion section. 14 Perchloric acid-potassium hydroxide Recoveries of C-PAH 14 and C-aPAH were evaluated after H^ClG^ deproteinization and neutrali zation with KOH by analyzing triplicate samples prepared as outlined in 14 Table ?.. The C-PAH standard was an aqueous solution containing 14 approximately 1.5 yCi in 100 yg of PAH per ml. The C-aPAH standard was an aqueous solution containing approximately 0.75 yCi in 200 jjg of aPAH per ml. 13
Table 2. Treatments for evaluation of perchloric acid and potassium hydroxide
a \^atev Blood ^ " Treatment (ml) (ml) ^ml)^
Control/Control 1350 (C/C)
Control 850 - 500 (C)
Blood 350 500 500 (B)
3 14 TrijJicate samples of each treatment contained 50 yl of C-PAH or C-aPAH standard solution.
Treatments were mixed by vortex stirring and centrifuged for 20
minutes at approximately 13,000 x gravity. Supernatants were filtered
through tissue paper and 1.5 ml transferred to another centrifuge tube.
This tube was placed on ice for 10 minutes after which 500 yl of 2.0 M
KOH was added and mixed well. After an additional 10 minutes on ice and
centrifugation for 20 minutes at approximately 13,000 x gravity, the
supernatants were decanted into sample vials. A 25 yl aliquot of the
fluid was injected onto the HPLC and the eluate collected as the PAH or
aPAH absorbance peak occurred.
The collected eluates were counted on a Beckman 8000 liquid scintilla tion spectrophotometer after the addition of 15 ml of Beckman Ready-Solv scintillation fluid. The observed CPM were converted to DPM by effi 14
ciencies calculated with the respective "H" numbers. The DPM values were
averaged within each sample and the percentage recoveries calculated and
summarized In the Results and Discussion.
Peak-height evaluation To evaluate peak-height linearity and peak-height ratio constancy, H^ClO^-KOH supernatants from whole blood
samples containing varied amounts of PAH, aPAH, and PAB were injected onto
the HPLC system. To 5 ml of whole blood, 2.5 ml of internal standard were added. The internal standard was an aqueous solution of PAH, aPAH, and
PAB in concentrations ranging from 0.12 to 64.0 pmoles of each compound
per ml. Complete hemolysis was assured with mixing, and 2.5 ml of 1 M
HgClO^ was slowly added while vortex stirring. After a minimum of 10
minutes, on ice, samples were centrifuged for 15 minutes at approximately
10,000 X gravity, and supernatants were filtered through tissue paper into
another tube. Cooling the sample after adding H^CIO^ was not absolutely
necessary, although better separation of the fluid and precipitated
protein occurred if the sample was placed on ice. A 1 ml volume of 2 M
KOH was added and the mixture cooled on ice for a minimum of 10 minutes.
After centrifugation for 15 minutes at about 10,000 x gravity, supernatants
were decanted from the firm potassium perchlorate pellet into a sample
vial and then frozen until analyzed in triplicate by HPLC as 25 yl aliquots.
Peak heights were measured for the three compounds, PAH:PAB and aPAHrPAB peak-height ratios calculated, and reported in Results and
Discussion. 15
Results and Discussion
Detection wavelength
The ultraviolet absorbance spectra for PAH, aPAH, and PAB are shown
In Figure 2. All three compounds exhibit absorbance maxima between 260
and 280 nm. Detection In this range would provide maximum detection.
In previously reported HPLC procedures that use UV absorbance detection
(Brown, et al., 1974; Brown, et al., 1976; and Brown & Mlchalski, 1976),
absorbance was monitored at 254 nm. The absorbance spectra in Figure 2
clearly show that absorbances at 254 nm are less than maximal; rather, a
steep slope occurs. Detection at 270 nm provides for greater sensitivity
and less variability in absorbances that might be caused by minor
fluctuations in the emitted wavelength from the monochrometer.
Electrochemical oxidation detection, reported by Shoup and Kissinger
(1975), provides good sensitivity for PAH detection, but requires a
specialized detector. Their report made no mention of detection and
quantitation of aPAH; it is unclear if their system is capable of detecting
aPAH, using the same settings used for PAH. It is also unclear if their
system allows for the use of and detection of an Internal standard such
as PAB.
Chromatographic conditions
Figure 3 is a chromatogram of a standard aqueous solution containing
the three compounds. The baseline resolution and peak symmetry allows
use of peak height measurements for quantitation, although peak areas 16
— PAH — PAB — aPAH
LU 3.00 O 1 g o CO 2.00
1.00
0.00 200 225 250 275 300 WAVELENGTH (nm)
Figure 2. Absorbance spectra of PAH, aPAH, and PAB The 1 mM solutions were prepared to pH 3.5 as described in the Materials and Methods. 17
PAB
PAH
z i 8 aPAh
V ir ± 12 3 MINUTES
Figure 3. Chromatograph of PAH, aPAH, and PAB aqueous standards containing 10 yg of each compound per ml 18
probably would provide more precise results. Inclusion of the PAB internal standard minimizes errors that may be introduced by non-quanti tative transfers of supernatants during sample preparation. Quantitation of PAH and aPAH then can be accomplished by peak-height ratios; i.e., comparison of the respective peak heights to the peak height of PAB within the same sample. Appendix A contains the procedure using a PAB internal standard for quantitation of unknown concentrations of PAH or aPAH.
Detector Linearity
The plot of detector response (in mm peak height) versus amounts injected onto the column (in nmoles) and the least-squares estimate of the linear equations appear in Figure 4. Correlation coefficients of 0.998 or greater obtained for each line indicate a strong linear relationship between peak heights and amounts injected (Snedecor & Cochran, 1967).
It can be concluded that a linear relationship exists between recorder response and amount of compound.
Deproteinizatlon
Any HPLC analysis of whole blood or plasma samples requires that protein be diluted or removed prior to injection onto the system. A concentration of 3 to 1600 nmoles per 25 pi aliquot corresponds to 0.12 to 64 ymoles per ml, or 2.8 to 1518 mg per ml for PAH and 2.4 to 1295 mg per 100 ml for aPAH. Clearance studies conducted by Smith, et al. (1945) 150
100
0 200 400 600 800 1000 1200 1400 1600 NMOLES INJECTED ONTO COLUMN
Detector response plot for PAH, aPAH, and PAB Duplicate solutions were prepared as described in the text and injected in triplicate as 25 yl aliquots. 20
utilized minimum PAH and aPAH concentrations of 1.5 and 3 mg per 100 ml, respectively, which are used as estimates for minimal plasma concen
trations for PAH and aPAH in blood-flow determinations. If deproteiniza- tion is not done, a dilution of 100 times or more is needed to dilute the
protein contained in blood prior to HPLC analysis. Therefore, dilution of samples is unsuitable for PAH and aPAH determinations because the concentrations of PAH and aPAH would be below the detection limits of the
HPLC system. A deproteinization step is necessary for HPLC analysis of
PAH and aPAH from whole blood.
In recovery experiments used to evaluate barium hydroxide-zinc sulfate and perchloric acid-potassium hydroxide methods, the comparison of "control/control" recovery of radioactivity with "control" and "blood" treatments and of "control" with "blood" treatments would reveal any loss of PAH or aPAH attributable to the deproteinization procedures. (See
Table 1, p. 11, and Table 2, p.13, for explanation of treatments.) A loss of radioactivity would occur in the "control" treatment as compared to the "control-control" treatment if the deproteinizing agent alone caused a co-precipitation of either compound. The only difference between the
"control/control" treatment and the "control" treatment was in the presence or absence of deproteinizing agent. Loss of radioactivity would occur between the "control" and "blood" treatments if the removal of protein caused a co-precipitation of either compound; the only difference between these two treatments is the presence ("blood" treatment) or absence
("control" treatment) of whole blood. 21
Barium hydroxide-zinc sulfate Bush, et al. (1979) accomplished
deproteinization of blood for analysis of volatile fatty acids (VFA) by
HPLC by the method of Somogyi (1945). Because VFA will be quantitated
during trans-liver balance studies in which PAH will be used as a blood-
flow marker, it would have been ideal if the same sample could be used
for determination of the blood-flow marker concentration as well as for
VFA concentration. 14 Results from C-PAH recovery experiments, summarized in Table 3,
indicate that approximately 25% of the is lost when Ba(0H)2 and
ZnSO^ is present ("control" and "blood") or absent ("control/control").
14 a Table 3. Percentage recoveries of C following Ba^OH^g-ZnSO^ treatments
Comparison Calculation^ Recovery
C versus C/C 2530/3365 x 100 75.2%
B versus C/C 2507/3365 x 100 74.5%
B versus C 2507/2530 x 100 99.1%
^Treatments and abbreviations are discussed in the preceding Methods and in Table 1, page 11.
^DPM values presented here are based on averages obtained from duplicate samples, and are associated with a standard error no greater than 2%.
This indicates that the precipitation of BaSO^ alone is responsible for the loss of C-PAH. A 99% recovery of ^^C in the "blood" treatment in comparison to the "control" treatment also indicates protein had little 22
effect on co-precipitation of C-PAH. Probably, the Ba(OH)2-ZnSO^ deproteinization effects occur by reactions with the glycine side-chain of PAH. Ohara and Ariyoshi (1979) reported a similar degree of amino acid loss when Ba(0H)2-ZnS0^ were used for deproteinization. Because of the loss of ^^C-PAH, the BaCOH^g-ZnSO^ deproteinization method was abandoned. 14 Perchloric acid-potassium hydroxide The recovery of C-PAH following perchloric acid deproteinization and neutralization by KOH is presented in Table 4. No significant differences in recovery were found
14 Table 4. Percentage recovery of C-PAH after perchloric acid-potassium hydroxide treatments
Comparison Calculation^ Recovery
c versus C/C 4243/4290 X 100 98.9%
B versus C/C 4294/4290 X 100 100.1%
B versus c 2494/4243 X 100 101.2%
^Treatments and abbreviations are presented in the preceding Methods and in Table 2, page 13.
^DPM values used here are based on averages from duplicate samples, and are associated with standard errors no greater than 2%. among the three treatments. The same comparisons as those used in evaluating the Ba(OH)^-ZnSO^ deproteinization method were 23
considered for perchloric acid-potassium hydroxide. Therefore, it was concluded that (1) neither perchloric acid nor the precipitation of potassium perchlorate caused a loss of ^^C-PAH, and (2) the precipitation of protein by perchloric acid did not cause an appreciable loss of ^^C-PAH.
Potassium hydroxide is included in the deproteinization procedure because the perchlorate ion is incompatible with the sialic acid support 18 resin of the C HPLC column. The potassium ion from KOH combines with the perchlorate ion to form insoluble potassium perchlorate, which precipitates. The volume of KOH added is minimized by use of a relatively high concentration so that little dilution of the sample occurs. Super- natants from blood samples have a pH above 10 after the addition of KOH to cause potassium perchlorate precipitation. Due to the small injection volume relative to the mobile phase buffer, however, the pH of the injected aliquot presumably is quickly decreased to 3.5 for separation of 18 the three compounds on the C column. Addition of concentrated H^PO^ may be necessary, however, to assure adequate separation. 14 The recovery of C-aPAH following perchloric acid deproteinization and neutralization with KOH is presented in Table 5. The presence or 14 absence of H^CIO^ or whole blood had little effect on the recovery of C radioactivity, although aPAH tended to be slightly affected by the precipitation of protein as revealed by the 96.9% recovery in the "blood" treatment versus "control" treatment. There is also a slight depression in recovery when the "blood" treatment is compared to the "control/control".
A loss of radioactivity in the "blood" treatment could be a result of 24
14 Table 5. Percentage recovery of C-aPAH after perchloric acid-potassium hydroxide treatment
Comparison Calculation^ Recovery
C versus C/C 2262/2220 x 100 101.9%
B versus C/C 2193/2220 x 100 98.8%
B versus C 2193/2262 x 100 96.9%
^Treatments and abbreviations are presented in the preceding Methods and in Table 2, page 13.
^DPM values reported here are based on averages from duplicate samples, and are associated with standard errors no greater than 2%.
14 CO-précipitâtion of C-aPAH with protein, a transformation of the aPAH molecule to one that does not elute during the aPAH absorbance peak, or a decrease in the detection of radioactivity in blood filtrates. It is unlikely that the H^ClO^-KOH treatment caused a chemical transformation, however, because of the recovery of 101.9% of the radioactivity in the
"control" versus the "control/control" treatments. This comparison evaluates the role that the deproteinizing agent alone plays in a loss of 14 C-aPAH as these two treatments differ only in the presence of perchloric acid and KOH, It is unlikely that detection of radioactivity was a problem because of CPM conversion to DPM by use of counting efficiencies. 25
Perchloric acld-KOH treatment does not affect the quality of
chromatography. A chromatogram from a supernatant of whole blood
deproteinized with perchloric acid and neutralized with KOH is illus
trated in Figure 5. This blood was taken from a steer injected with PAH
and aPAH 20 minutes prior to collection of the blood sample. Comparison
of Figure 5 with Figure 3 (p. 17) illustrates that no resolution or
symmetry of peaks is lost by the deproteinization procedure.
It is critical that no PAH or aPAH be removed from samples during
preparation for the HPLC analysis. This is also true for the internal
standard, PAB, which is added to the samples Immediately after collection
from the animal. In order that the PAB internal standard be used with
confidence, the peak-height ratios of PAH:PAB and aPAHrPAB must be constant
in the range of concentrations to be encountered during blood flow
experiments. Table 6 contains the peak-height ratios for both combinations.
The ratios remain relatively constant over a wide range of concentrations. 14 No C-PAB was available for recovery experiments that were used to
evaluate possible losses of PAH and aPAH. However, because the peak-
height ratios were constant over the wide concentration range, obviously no appreciable loss of PAB occurred. The function of the Internal
standard solution is twofold: 1) to provide a vehicle for addition of
internal standard, and 2) to provide a hypotonic medium so that complete
hemolysis occurs prior to deproteinization. 26
PAH
S s
J L 012 3 MINUTES
Figure 5. Chromatogram of a perchloric acid-KOH filtrate of whole blood collected from a steer injected with PAH and aPAH 20 minutes prior to sampling. 27
Table 6. Peak-height ratios for PAHtPAB and aPAHtPAB obtained from whole blood after perchloric acid-potassium hydroxide treatment
Amount injected PAH:PAB aPAH:PAB (nmoles) ratios ratios
20 1.45 (0.06) 0.98 (0.03) 40 1.46 (0.02) 1.00 (0.04) 100 1.61 (0.01) 1.12 (0.02) 200 1.53 (0.01) 1.21 (0.01) 400 1.58 (0.01) 1.12 (0.01) 600 1.57 (0.01) 1.11 (0.01) 800 1.58 (0.004) 1.12 (0.006) 1200 1.54 (0.002) 1.12 (0.002) 1400 1.54 (0.003) 1.12 (0.003) 1600 1.55 (0.004) 1.13 (0.004) X = 1.54 (0.05) 1.10 (0.07)
®The ratios are averages obtained from 25 pi aliquots injected in triplicate. The standard error appears in parentheses.
Conclusions
Several i^nportant conclusions can be reached from the evidence of
Part I: 18 1. An HPLC procedure that utilizes a reverse-phase C column and
a flow rate of 2.5 ml/minute of a mobile phase of 30% methanol
(V/V) in 10 mM NaH^PO^ buffer, pH 3.5, provides for an excellent
separation of PAH, aPAH, and PAB in approximately 3 minutes.
2. No appreciable loss of either PAH or aPAH occurs during
deproteinization with perchloric acid and subsequent neutral
ization with KOH. 28
3. Resolution and symmetry of the PAH, aPAH, and PAB peaks are not
affected by perchloric acid deproteinization followed by
neutralization with KOH.
4. Peak-height ratios of PAH:PAB and aPAHrPAB remain constant over
a wide range of concentrations. This range is expected to
include concentrations to be encountered during studies using
PAH for determination of blood flow.
Development of this HPLC procedure provides a rapid and accurate method for quantitation of PAH and aPAH. A significant improvement over previously available methods is provided by this system because no coupling reactions are needed for detection, which is accomplished by direct UV absorbance. Also, accuracy of aPAH analysis is greatly improved because no acid hydrolysis is required; again, the detection is by direct means. The detection system also is more sensitive at 270 nm for the three compounds than at the 254 nm setting used for some previously reported HPLC methods. 29
PART II. PARA-AMINOHIPPURIC ACID
IN VIVO
Literature Review
The Literature Review of Part II will present some brief information
on several techniques for measurement of blood flow in animals and a more
detailed discussion of dye-dilution techniques, especially in regard to
PAH as a blood-flow marker. Although PAH has been used as a blood-flow
marker, the vast majority of work using PAH has been concerned with
elucidation of organic anion transport mechanisms in kidneys. No attempt will be made to present an exhaustive review of all that is known about
the transport mechanism of PAH. Rather, a brief discussion of the renal
transport will be given. What is known about the metabolism of PAH will
be discussed In detail.
Some techniques for the determination of blood flow
This major section will briefly cover many of the techniques used for
blood flow with the exception of Indicator-dilution, which will be covered in detail In the next major section. Each technique will be described with its principle of operation as well as Its major disadvantages for Its application to trans-organ balance studies. General principles of blood flow measurement have been discussed in detail in a book by Woodcock (1975).
Electromagnetic flowmeters Electromagnetic flowmeters were developed in the 1930s (Kolln, 1936; Wetterer, 1937), and have evolved into many designs for a variety of applications (Clark & Wyatt, 1969). 30
The principle behind electromagnetic flowmeters is the induction of
voltage at right angles to the direction of fluid flow and an applied
magnetic field when the conducting fluid passes through the magnetic
field. Major disadvantages with these flow determinations have caused limited application in large animals. Variations in magnetic field strength causes significant changes in flow detection sensitivity
(Wyatt, 1961). Precise calibration must be conducted for accurate flow determination; this is rather difficult to do, especially in larger vessels. Brungsting and TenHoor (1968) list five factors that contribute to errors in calibration: (.1) hemoglobin concentration of blood,
(2) probe temperature, (3) fit of the flowmeter around the vessel (diffi cult with large vessels), (4) insulation of the probe around the vessel, and (5) axial symmetry of the velocity of flow within the vessel. Some doubts also exist regarding the safety of electromagnetic flowmeter probes in vivo because some probes generate significant amounts of heat
(Jones & Wyatt, 1970). These flowmeters require specialized electronic equipment for interpretation and manipulation of the transducer output.
Ultrasonic flowmeters Two general types of ultrasonic flowmeters have been developed in the last 20 years: (1) those that measure time-of- arrival differences between ultrasound emitted with and against the direction of blood flow, and (2) those that monitor the change of frequency, or Doppler-shift, of emitted ultrasound. In instruments of the first type, the difference in phase between the emitted wave travelling opposite to the fluid is proportional to flow of fluid. The second type 31
exploits the fact that the frequency shift of the scattered high-frequency
emitted sound wave is proportional to the velocity of the scattering
particles, which are the red blood cells. Both principles are discussed
in detail in Chapter 5 of Woodcock (1975). The diffraction of emitted
wavelength and the physics of the scattering process are important con
siderations in ultrasonic flowmeter use.
Woodcock (1975) also discusses important factors that may affect the
ultrasonic determination of blood flow: (1) hematocrit, (2) amplitude and
phase distortion of the recorder system, and (3) axial drift of blood
cells in the measured blood vessels. Also, the signal-to-noise ratio of
the ultrasonic detector system is decreased as the cross section of the
vessel being measured is increased (Gosling, et al., 1970). When deter
mining blood flow in large animals such as cattle, vessel diameters are
large enough that the signal-to-noise ratio may significantly decrease the sensitivity of flow measurements. Ultrasonic systems, like electro magnetic flowmeters, require expensive electronic equipment for data collection and interpretation. Other problems associated with ultrasound also are discussed by Woodcock (1975), but a lengthy discussion is beyond the scope of the purpose of this Literature Review.
Thermal methods Some of the earliest techniques developed to estimate blood flow involved some type of measurement involving heat or temperature: (1) calorimetry (Stewart, 1911-1912), (2) thermal dilution
(Lochner, 1953), (3) resistance thermometry, (4) thermal conductivity changes (Gibbs, 1933), and (5) skin temperature as an indication of blood 32
flow (Eddy & Taylor, 1931). Thermal methods have disadvantages in that
they require constant attention for calibration and correction by frequent
measure of systemic blood temperature, that hemoglobin can alter the heat
capacity, that the exact amount of thermal indicator injected is difficult
to measure and control, and that hemodilution is a problem in long-term experiments. Thermal techniques also are too complicated and lack the sensitivity that is required in trans-organ balance studies.
Other methods Pressure-sensitive flowmeters determine blood flow by measuring the frictional decrease of pressure. According to Woodcock
(19 75), three parameters can be responsible for differences in pressure between two points within a tube; (1) local acceleration of fluid (this changes only when flow rate changes with time), (2) convective acceleration caused by a change in the cross-sectional area of the flow tube or localized stagnation of flow: i.e., flow eddies, and (3) frictional loss of pressure. Several types of pressure-sensing probes and devices have been developed. Pressure-change flowmeters, however, require several physical and mathematical assumptions which severely limit their application
(Woodcock, 1975).
Several types of outflow recording systems that measure flow by collection of venous blood were described by Green (1948). Several adaptations of collection methods have been made, among which are the earliest techniques for direct measurement of blood flow; e.g., Volkmann
(1850). One type of device, the Ludwig strohmuhr, collects blood from a cannulated vein for a known period of time and then returns blood to the 33
systemic circulation after volume is measured. In bubble-flowmeters, the
flow of blood in the blood vessel is directed through an external tube
where a bubble is introduced. The length of time required for the bubble
to travel the known distance of tubing is used to calculate the flow of
blood. These collection methods suffer primarily from cannulation problems
and the requirement of anti-coagulants (Woodcock, 1975).
Nuclear magnetic resonance (NMR) first was used to measure blood flow
by Boifman and Kudravcev (1959). They utilized the NMR spectrum for
hydrogen for their calculation of blood flow. Due to the major expense
for NMR and problems associated with calibration and design for probes,
NMR techniques offer little, if any, advantage over other methods of
blood flow determination (Woodcock, 1975).
Magnetorheography is an adaptation of the electromagnetic principle.
The magnet and detecting electrodes are external and the induced electro
motive force is detected through a limb. The equipment required is costly and artifacts from muscular movement such as breathing are large. Also, this method cannot measure accurately the flow of blood across an organ; it was developed to estimate flow in appendages.
Aisenburg (1970) reported a flow method based on the distortion of an electromagnetic field pattern. The geometry of placement of the flow probe is critical and the sensing electrode must remain in the middle of the stream of flow. Again, this method would not be acceptable for calculation of blood flow in trans-organ balance studies. 34
General limitations The methods described to this point are
impractical or impossible for trans-organ balance studies for one or more
of the following reasons: (1) the method or equipment is too costly,
(2) there is a lack of sensitivity and/or accuracy, (3) placement of
probes or detectors situ or in conscious animals is impossible due to anatomical limitations, (4) the methods are too time consuming or complicated, (5) there is a lack of spatial or time relationship between sampling of blood for metabolite determination and determination of blood flow, and (6) the blood flow method itself may alter the flow of blood.
Determination of blood flow by indicator-dilution
A number of indicators have been used to estimate blood flow; e.g., heat, dyes, inert gases, and radioactively-labeled markers. The choice of indicator depends on the type of experiment and the characteristics of the indicator required for the blood flow determination. The discussion in this section will be limited to exogenous dyes as indicators because they are well-suited for trans-organ balance studies, although the principles for blood flow calculations remain the same regardless of the indicator.
The first injection of dye into blood for flow determination purposes is attributed to Gross and Mittermeyer (1926). The theory of dye-dilution determinations of blood flow has been presented extensively in discussions by Meier and Zierler (1954), Zierler (1962), Woodcock (1975), and Lassen and Perl (1979). The principle involves an injection of a known quantity of dye into the blood system as a single-injection (bolus) or as a 35
continuous infusion. Downstream from the injection site, blood is with
drawn at various time intervals and analyzed for concentration of dye.
From the amount of dye injected and the concentration of dye in blood
taken at the sampling site, flow versus time can be calculated.
Woodcock (1975) summarizes some assumptions made in indicator
dilution methods: (1) the indicator must have the same distribution
and transit times as blood, (2) the indicator must not recirculate (this
can be corrected for), and C3) the indicator in the system is eventually
eliminated. Meier and Zierler (1954) and Zierler (1962) also state that
flow in the system to be measured must be constant, and the system have
a single inflow and outflow vessel.
The dye-dilution method is the method of choice for blood-flow
determinations in trans-organ balance studies because of two important
considerations. First, in continuous infusions, after the infusion pump
is started and a blood equilibrium of indicator is obtained, the deter
mination of blood flow requires blood samples only. The samples can be
collected and frozen until a more convenient time for analysis. This
delay could provide valuable time during trans-organ experiments and
provide for more accurate timing of experimental procedures. The blood
flow dye and metabolite concentration in blood can be determined from
the same blood sample or from two closely-timed blood samples. Thus,
more accurate metabolite fluxes are calculated than those obtained from
less closely-timed determinations. Second, the determination of trans- liver flow is not dependent upon elaborate blood flow equipment that is 36
presently unavailable in the Nutritional Physiology Laboratories at Iowa
State University. Also, determination of flow does not depend upon
equipment that must be in good working order at the time of the experiment.
The flow data need not be obtained at the time of the experiment when dye-
dilution methods are used.
Several dyes have been used to determine flow of blood through the
liver. Indocyanine green (Cardiogreen) has been used extensively because
of its removal by the liver (Cherrick, et al., 1960; Fox & Wood, 1960;
Wangsness, 1971; and Paumgartner, et al., 1970). Because of its removal
by the liver, Cardiogreen can be used for determination of portal vein
blood flow, but it would be of no use for simultaneous hepatic venous flow
determinations. Bromosulfophthalein (BSP) has been used for hepatic venous flow since its introduction by Bradley, et al. (1945). However, neither of these compounds alone can be used for simultaneous determination of portal and venous flows in and out of the liver.
It is of importance that simultaneous blood flows of the hepatic vessels be determined for trans-liver experiments so that accurate metabolic fluxes can be determined. Therefore, the method developed by
Katz and Bergman (1969a) seemed an attractive candidate for the trans-liver experiments. Their method, an extension of one by Roe, et al. (1966),
used PAH. However, their method was reported for use in sheep and dog;
little information if available about the use of PAH in cattle. Mangelson,
et al. (1968) reported the clearance of PAH to be approximately 1200 2 ml/(min x m body surface area) in dairy cattle. 37
Renal transport of PAH
PAH and hippuran, the ortho-iodo derivative of hippuric acid,
originally were considered as indicators for the evaluation of renal
plasma flow and clearance (Elsom, et al., 1936; and Smith, et al., 1938).
Because it was demonstrated by Cross and Taggart (1950) that rabbit
kidney was capable of concentrating PAH in cortical slices, however, PAH
has been used extensively to investigate the transport of organic anions
in renal tissue.
A generalized model of PAH transport in renal cortical tissue has
been described by a number of authors (Chinard, 1956; Foulkes & Miller,
1959a and 1959b; Tune, et al., 1969; and Kinsella, et al., 1979). In
general, the steps of PAH transport are believed to occur in the following sequence: (1) diffusion of PAH from blood plasma to cortical extra-
vascular space, (2) active transport of PAH from the extravascular space at the peritubular cell membrane, (3) concentration of PAH within the peritubular cell, and (4) passive diffusion of the exchangeable fraction of intracellular PAH down a concentration gradient into the tubular lumen.
Apparently only the antiluminal membrane of the lumen cell is capable of concentrating PAH. Diffusion of PAH from the cell to the tubular lumen is passive and does not seem to be mediated by a membrane carrier
(Kinsella, et al., 1979). Also, two intracellular pools of PAH evidently exist; one which is bound or compartmentalized and equilibrates slowly with a second pool that is freely-diffusable (Welch & Bush, 1970).
The PAH transport system can be inhibited by metabolic poisons: two 38
key ones are 2,4-dlnitrophenol (DNP) (Mudpe & Taggart, 1950a; Copen-
haver & Forster, 1958; Huang & Lin, 1965; Shideman, et al., 1952;
and Eveloff, et al., 1979) and ouabain (Podevin & Boumendil-Podevin,
1977; Podevin, et al., 1978; Holliday & Miller, 1980; Spencer, et al.,
1979). A number of other inhibitors have been identified; however, a
detailed discussion of various inhibitors will not be presented. In
general, the inhibitors fall into two categories; first, those that
inhibit PAH transport by competitive binding to the transport mechanism,
and second, those that interfere with the active uptake mechanism.
Several stimulators of PAH uptake also have been described. 3 Autoradiographic studies using H-PAH have shown that intracellular
accumulations are evenly distributed throughout the cytoplasm and
excluded from the nucleus (Miatello, et al., 196 6; Wedeen and Weiner, 1973a,
and 1973b; Weinman, et al., 1978; Woodhall, et al., 1978). Uptake and secretion is limited to the straight portion of the proximal tubule
corresponding to the pars recta (Voodhall, et al., 1978; Weinman, et al.,
1978). Little difference was noted between superficial medullary and juxtacortical proximal tubules.
Because the location and characteristics of PAH transport are so well understood, PAH transport has been used widely as a standard for renal anion transport to which the transport of other compounds is compared
(e.g., Schali & Roch-Ramel, 1980; Roch-Ramel, et al., 1980; Hong, et al.,
1978; Bito & Baroody, 1978; Barac-Nieto, 1971; Kuo & Hook, 1979).
Also, PAH transport has been used as a standard to evaluate the develop ment and maturation of the "renal organic anion" transport mechanism in 39
neonates (Bond et al., 1976; Hewitt & Hook, 1978; Kluwe, et al., 1978;
and Noordewier & Withrow, 1979).
PAH clearances and kinetics in vivo
The renal clearance of PAH has been calculated for a variety of conditions in dogs (Smith, et al., 1945; Pritchard, et al., 1965;
Blaufox, et al., 1963; Bond, et al., 1976; McDonald & Kelley, 1948), humans (Smith, et al., 1945; Newman, et al., 1949: Schnurr, et al., 1980), rats (Sadjak, et al., 1979; Chiu & Long, 1979), sheep (McDonald & Mac-
Farlane, 1958) and pigs (Gyrd-Hansen & Rasmussen, 1970). The values range from 16 ml/min for the puppy (Bond, et al., 1976) to approximately
700 ml/min in the human (Newman, et al., 1949).
In the dog and human, PAH and aPAH are handled similarly because there are identical clearances for PAH and for aPAH (Smith, et al., 1945;
Gyrd-Hanson & Rasmussen, 1970). In the human, clearance decreases for
PAH and aPAH with decreasing concentrations, whereas clearances remain the same in the dog regardless of plasma concentrations of PAH and aPAH
(Newman, et al., 1949). Haberle, et al. (1978) state that there is a correlation of the amount of PAH secreted with the glomerular filtration rate, but no relation exists between PAH secretion and PAH plasma "load" or renal plasma flow. Girndt, et al. (1974b) observed faster clearance for aPAH than for PAH, as revealed by finding more N-acetyl-PAH in the urine than in mixed venous blood. Gyrd-Hansen and Rasmussen (1970) reported an aPAH:PAH clearance ratio of 1,08. 40
Hippuran has been used extensively to investigate the disappearance
of hippuric acid derivatives from blood, although very, little information is available for PAH in this regard. However, information obtained with hippuran may relate closely to PAH because hippuran and PAH have identical excretion and clearance rates (Smith, et al., 1945).
Hippuran was used first by Smith, et al., (1938) to determine tubular excretory mass, effective blood flow, and filtration rate. Also, 131 I-hippuran has become a useful tool in kinetic investigations due to the ease of its analysis (Magnusson, 1962; Pritchard, et al., 1965;
Blaufox, et al., 1963). When renal blood flow determined by direct measurement (collection of blood) was compared with flow determined by a 131 I-hippuran single-injection method, no difference was obtained 131 (Pritchard, et al., 1965). I-hippuran was used by Blaufox, et al. 131 (1963) simultaneously with PAH to evaluate use of I-hippuran for renal function tests. They found no difference between disappearances of the two compounds. When Maher and Evleback (1970) simultaneously injected 125 131 I- and I-orthoiodohippurate with PAH, no differences were noted among clearances of the three compounds. Similar results were obtained when unbound PAH concentrations were used to measure clearance (Maher, et al,, 1971).
Mandel, et al. (1955) investigated the disappearance of PAH following single injections in dogs. They found a PAH disappearance curve from blood that could be resolved into two components when plotted on a sémilogarithmic (base e) scale. The "rapid" component was described by a 41
line that had a Y-lntercept of 0.99 and a slope of -0.291. The "slow"
component was represented by a line having a Y-intercept of 0.48 and a
slope of -0.0375. A three-compartment model was used by VanStekelenburg,
et al. (1976) to describe the disappearance of PAH from human blood.
In comments regarding the VanStekelenburg model, Nowotny and Kletter
(1976) mentioned that the component having the most rapid disappearance
was due to the outflow pattern of the lower extremity rather than to a
general body compartment. That is, the most rapid compartment could be
described merely by the blood flow pattern of the leg rather than an
overall distribution of hippuran in three compartments in the body in
general. Therefore, there is some doubt about the validity of a three-
component over a two-component model. Also, no report is known that treats
the compartmental analysis in terms of PAH and aPAH interconversion.
Previous models are involved with physical transport kinetics.
A physiological space that corresponds to the first, and most rapid, component of disappearance curves has been difficult to describe. Several authors have indicated various degrees of plasma protein binding of PAH 131 or hippuran. Magnusson (1962) used I-hippuran to evaluate protein binding by protein precipitation, dialysis, ultrafiltration, and electro phoresis. He found that approximately 1% of the radioactivity precipitated with protein, that about 1.2% was undialyzable, that about 4% was unfilterable after 20 hours, and that unbound PAH migrated faster than serum proteins during electrophoresis. He concluded that either none or very small amounts of radio-active hippuran combined with serum proteins. 42
Maher, et al. (1971) Investigated the extent of plasma binding by ultra filtration techniques and found that approximately 33% of o-iodohippurate and 74% of PAH was free in venous plasma. Bond, et al. (1976) found approximately 40% of ^^C-PAH was undialyzable in vitro after 24 hours at at 39°C when 4, 12, or 20 mg/100 ml PAH concentrations were present in plasma. Mudge and Taggart (1950a)found PAH to be freely dialyzable.
Ultrafiltration techniques have shown that approximately 44% of the PAH found in rabbit blood seemed to be bound to plasma proteins (Foulkes, 1963).
Rahill (1970) found approximately 17% of the PAH bound at 37° by dialysis.
It is not known if protein binding contributes significantly to rapid disappearance of PAH from blood in cattle.
It is possible also that rapid disappearance of PAH in vivo can be explained in part by PAH being sequestered by red blood cells. Smith, et al. (1945) found no penetration of PAH into human erythrocytes even after prolonged infusions, although other evidence indicated some pene tration in dog erythrocytes. Magnusson (1962) found with rats that radio activity of radioactive hippuran diminished at the same rate in plasma and whole blood, even though about 40% of the total radioactivity of whole blood was recovered in red blood cells. He concluded that even if
PAH does enter red blood cells, the membrane must be adequately permeable to allow free exchange with plasma. VanStekelenburg, et al. (1976) postulated that the contribution of red blood cell uptake to the dis appearance might be of significance to the second component of their three- component model of hippuran distribution. Girndt, et al. (1974a) state 43
that entrance of PAH into erythrocytes of dogs occurs at a rate that would
not contribute to errors in clearance calculations. It is not known if
uptake of PAH into blood cells contributes to a significant fraction of
PAH in whole blood in cattle.
Metabolism of PAH
PAH is conjugated to its N-acetyl derivative in several species:
human, rabbit, pig, guinea pig, mouse, cat, cattle, sheep, and rat
(Malyusz, et al., 1979; Carpenter & Mudge, 1980; Smith, et al., 1945;
Despopoulos, 1956; Frindt & Vial, 1968; Setchell & Blanch, 1961;
Gyrd-Hansen & Rasmussen, 1970; Cross & Taggart, 1950; Girndt, et al.,
1974a and 1974b), but not in dogs (Mudge & Taggart, 1950b; Newman, et al., 1949). Of these reports, only Setchell and Blanch (1961) indicate a conversion of PAH to aPAH in kidney slices from a cow. They calculated average conjugation rates of 0.54 and 0.15 ymole/g wet weight/hr for kidney and liver slice, respectively. No significant conjugation was noted with slices of spleen, lung, or heart muscle. However, no known report provides conversion rates of PAH for cattle in vivo.
Conversion of PAH to aPAH seems to be dependent upon a number of influences. Carpenter and Mudge (1980) reported wide variability in acetylation rates among four strains of mice and found that male mice had conversion rates four times that of females. With rats, Malyusz, et al. (1979) found that an insufficient supply of oxygen reduces the conversion of PAH, presumably by changing the tissue pattern of non es terified fatty acids, which in turn inhibits the N-acetylating enzymes. 44
This indicates that the availability of oxygen influences acetylation rates. Gimdt, et al. (1974a) found decreased acetylation rates in man with severe essential hypertension and decreased renal flow. The decreased rate of acetylation caused by 2,4-DNP in mouse kidney slices and other evidence just cited support the theory that acetylation is dependent upon high-energy phosphate bonds (Carpenter & Mudge, 1980). Age does not seem to be an important factor of acetylation in pigs; a 50 to
70% conjugation rate of PAH to aPAH was constant over wide age differences
(Rennick, et al., 1961).
Transformation of PAH to aPAH occurs in the kidney, and to a lesser extent, in the liver (Setchell & Blanch, 1961; Despopoulos, 1956; Frindt
& Vial, 1968). However, a possibility exists that some metabolism occurs in parenchymal tissue of other organs (Girndt, et al., 1974b) although it has not been quantitatively determined and reported.
De-acetylation of aPAH to PAH has not been found to occur in kidney slices in rabbits or rats (Newman, et al., 1949; Mudge & Taggart, 1950b;
Carpenter & Mudge, 1980); however, Girndt, et al. (1974b) discussed the possibility of extra-renal sites of de-acetylation. Metabolic products from PAH other than aPAH (e.g., N-acetyl-p-aminobenzoic acid) have been observed (reported by Frindt & Vial, 1968, and summarized by Girndt, et al., 1974a), although little Information is available. The potential errors introduced into clearance and flow calculations by the transformation of PAH is discussed in detail by Girndt, et al. (1974b).
Several points can be made in summary. First, PAH is actively trans 45
ported by renal peritubular cells at the antiluminal plasma membrane and
passively diffuses into the tubular lumen. Second, the kidney is
essentially the exclusive site at which PAH is removed from blood for
excretion. Third, PAH is acetylated to form aPAH primarily by the
kidney, and to a lesser extent by the liver. Little is known regarding
the in vivo acetylation in cattle. Fourth, de-acetylation of aPAH to PAH
does not occur in liver or kidney slices. It is not known if there are
extra-renal sites of de-acetylation. Fifth, conflicting reports exist regarding the extent PAH is bound to proteins (0 to 44% bound) or the extent PAH enters the red blood cells.
Materials and Methods
Experiments with non-radioactive PAH and aPAH
Single-injection and continuous-infusion experiments were performed with PAH and aPAH in an attempt to characterize the in vivo behavior of
PAH and aPAH. Steers were obtained from the Iowa State University dairy herd and were fed 1.15 kg of 70% chopped alfalfa hay and 30% concentrate
twice daily. The concentrate consisted of 92% corn, 5% monosodium
phosphate, and 3% trace mineral mix.
Sampling of blood was timed from the beginning of the single injection
or from the beginning of the continuous infusion and primer dose. Jugular
catheters were flushed with 2-3 ml of 0.9% saline containing 20 U
heparin/ml after each sample was collected.
Single injections A total of 27 experiments were conducted on
five steers weighing 80-115 kg. PAH was injected into all five steers
in 14 experiments; aPAH was injected into four of these steers in 13 46
experiments. One steer (#8229) developed an upper respiratory infection and was omitted from aPAH injections.
On the morning of experiments, a 5h" x 14 gauge Angiocath catheter
(Deseret Pharmaceutical Co., Sandy, Utah) was placed in a jugular vein and sutured to the skin. After a minimum of 1 hour, a control blood sample was withdrawn from the catheter, then a PAH or aPAH injection was administered during 1 minute. Dose rates were 15, 30, or 45 mg/lb
BW. The injection consisted of an appropriate volume of a 60 mg/ml sterile aqueous solution of PAH or aPAH, pH 7.3 prepared in 0.9% saline.
At 2, 5, 10, 15, 20, 30, 45, 60, 90, 120, 180, and 240 minutes after starting the injection, a 2-3 ml sample of blood was withdrawn and dis carded, then a 6 ml sample of blood was collected. To 5 ml of the blood,
2.5 ml of PAB internal standard (216 yg/ml) was added. After complete hemolysis was assured by vortex mixing, 5 ml of 1 M H^CIO^ was added. The mixture was centrifuged for 15 minutes at approximately 7,000 x gravity, and the supernatant was filtered into another tube through tissue paper.
A 1.25 ml volume of 2 M KOH was added, and the tube was placed in ice for a minimum of 10 minutes. The samples were centrifuged at approximately
7,000 X gravity for 15 minutes to sediment potassium perchlorate, and the supernatants were decanted into sample vials. These supernatants were stored frozen until analyzed by HPLC.
Concentrations of PAH and aPAH were determined by peak-height ratios as described in Appendix A. Adjusted concentration values were calculated by the following equation:
Adjusted Concentration = ^Ug/rol of compound^ ^ loo. yg total dose 47
Adjusted concentration values were used rather than actual concentration
so that the results would be standardized among all experiments. Values
were plotted on a log (base e) scale versus time, and the two component
slopes and Intercepts were calculated as described in the next paragraph.
Curve peeling, or graphical analysis, was performed to obtain the "Ic t slopes and Intercepts of a two component exponential function, Y » A^e 1
+ A^e ^2^. The first component, A^e ^1^, describes the rapid disappearance
from blood; the second component, A^e ^2^^ describes the slower component.
The observed values at times greater than 30 minutes are essentially free
from effects of the rapid component, and can be used to estimate values
for Ag and of the slower component. To calculate A2 (the intercept) and kg (the negative slope) of the slower component, base e logarithms of adjusted dose values for times greater than 30 minutes were analyzed by the general linear models (GLM) procedure of the SAS computation system (SAS
Institute, 1979). The antilog of the Intercept from GLM corresponds to the
Ag value; the slope is the negative value for kg. The estimates for A^ and kg were used in the equation, Yg = Age ^2^ (Yg=predicted value, t=time) to calculate the predicted adjusted concentration values for the slower component. The predicted values at each time, Yg, were subtracted from their corresponding observed values to obtain Yj, the adjusted concen tration that is removed by the rapid component. Natural logs of the Y^ values greater than 0 were analyzed by the GLM procedure to obtain estimates of A^, the Intercept, and k^, the negative slope, of the more rapid component. 48
Continuous infusions The same four steers weighing 145-160 kg were
selected at random for four experiments In which PAH was infused and for
five experiments in which aPAH was infused. The night before an experiment,
bilateral jugular catheters (Anglocath, 5h" x 14 gauge) were Inserted and
sutured to the skin. One-way valves were attached and sutured to the skin.
The catheter and valve were filled with heparlnized 0.9% saline. Valves
were capped, and injection sites and valves were covered with elastic tape
that went around the neck so that steers could not pull out the catheters
during the night.
A control blood sample was collected immediately before beginning the
infusion and primer injection. The priming injection of 5 mg/kg BW (given
as an appropriate volume of a 50 mg/ml sterile solution, pH 7.3, of PAH or
aPAH in 0.9% saline) and the continuous infusion were started simultaneous
ly by using both catheters. The continuous infusion was delivered by a
model 952 double-syringe infusion pump (Harvard Apparatus Co., Mlllls,
Mass.). The solution consisted of PAH or aPAH in a concentration of 0.25
mg/kg BW per ml, prepared sterile pH 7.3, In 0.9% saline, and was admin
istered at a rate of 1.0 ml/mln.
Blood samples were collected at 2, 5, 10, 20, 30, 45, 60, 75, 90, 105,
120, 150, 180, 210, and 240 minutes after the beginning of the primer and
infusion. The blood was processed in the same manner as described for the
single injections except that two samples were prepared at each time, and
1.5 ml of 60 yg/ml PAB Internal standard solution was added rather than 2.5
ml of 216 yg/ml used in the single Injeclons. Peak-height ratios with PAB were used to quantltate PAH and aPAH on the HPLC in the same way as for single injections, except that the sensitivity of the Omniscribe recorder 49
was ten times that for single injections.
Blood concentrations of PAH and aPAH were analyzed by the GIU pro
cedure (SAS Institute, 1979) using values from those samples collected
after 20 minutes to determine equilibrium concentration and slope. The
results of these experiments are discussed in the Results and Discussion.
Because of the high variability and lack of a consistent equilibrium
concentration among experiments, further experimentation was considered 3 necessary. Therefore, experiments utilizing H-PAH were performed.
Experiments with radioactive PAH
To overcome problems encountered during the investigations with non radioactive PAH just described, experiments utilizing a continuous infusion 3 of non-radioactive PAH and a single injection of (2-glycyl- H)-PAH
(Amersham, Arlington Hts., 111.) were performed. The rationale for this approach is discussed in detail in Results and Discussion.
Six Holsteln steers, weighing 70-110 kg, were obtained from the Iowa
State University dairy herd. They were fed a standard, calf-starter ration twice daily and had ad libitum access to water. The diet consisted of 40% com, 27% oats, 20% soybean meal, 10% molasses, 2% dlcalclum phosphate, 1% salt, and 0.1% of a calf premlx containing vitamins A and D. Experiments were conducted with steers in a metabolism crate with a urine collection pan. On the evening prior to an experiment, the skin overlying the jug ular veins was shaved, and body weight of the steer was recorded.
Bilateral jugular catheters (right side, Qulk-Cath, 2" x 16 gauge from
Vlcra, Dallas, Texas; left side, Anglocath, 5h" x 16 gauge or x 14
gauge from Deseret Pharmaceutical Co., Sandy, Utah) were inserted, sutured 50
to the skin, filled with 0.9% saline (20 U heparin/ml) and capped. The
steer was placed in the crate; and after approximately 1 hour, the "pre-
cold" control blood sample was taken, and a continuous Infusion of a
sterile aqueous solution containing 30 rag of PAH per ml at pH 7.3 was
begun. A double-syringe infusion pump (model 952, Hairvard Apparatus Co.,
Millis, Mass.) was used to deliver 2.5 ml of the infusate per minute
through the right jugular catheter.
Urine collection began when a steer was placed in the crate, and con tinued 3 hours past collection of the last blood sample. The time and volume of each urination were noted, and a sample of urine retained and frozen. 3 The H-PAH injection solution was prepared by adding approximately 1.5 3 9 ml of a H-PAH stock solution having an activity of 1.205 x 10 DPM/ml
(0.54 mCi/ral) to 25 ml of sterile 0.85% saline. The mixture was shaken well, and drawn into a 50 ml syringe. Three 5-ml washes of 0.85% saline were added to the syringe. An aliquot of injection solution contained in the syringe was taken so that total dose activity could be determined. The
"pre-hot" control blood sample was taken, after which the H-PAH solution was injected into the left jugular catheter during a period of 20 seconds.
Blood was withdrawn into the injection syringe and re-injected into the animal three times to assure total dose delivery. The time reference for 3 all samples and urinations was the beginning of the H-PAH injection.
Blood samples were collected via the left jugular catheter at 1, 2, 4,
6, 15, 20, 25, 30, 40, 50, 60, 75, 90, 105, and 120 minutes. For the first experiment, blood collection continued until 180 minutes; however, analysis of the blood samples revealed that radioactivity had decreased to 51
essentially background by 90 minutes and the third-hour samples were omitted from the remaining five experiments. Approximately 10 ml of blood was collected for all times except for 75 and 90 minutes, when 30 ml was taken so that adequate plasma was available for Evans blue determination. After the last blood sample, the infusion was discontinued, and both catheters removed. Steers remained in the crate for an additonal 3 hours for urine collection.
To 3 ml of blood, 1 ml of an aqueous solution of 1.28 M PAH and 1.28 M aPAH was added. After vortex mixing, 3 ml of 1 M H^CIO^ was added, and the tube was placed in ice. After centrifugation for 15 minutes at approx imately 7,000 X gravity, supernatants were filtered through tissue paper and 4.5 ml were transferred to another tube. A 0.5 ml volume of 3 M KOH was added, and the tube was placed in ice for at least 10 minutes. After again centrifuging for 15 minutes at approximately 7,000 x gravity, super natants were decanted into sample vials and frozen until analysis. To the remaining blood, 1 drop of 10,000 U heparin/ml was added before centri fuging to sediment the cells. Plasma was removed and frozen.
Hematocrits were determined at 15 minute intervals up to and including
90 minutes and including the two control blood samples. The packed-cell averages from all hematocrits were averaged and used in determination of whole blood volume from plasma volume.
Evans blue was used to determine plasma and whole-blood volume as out lined in Appendix B. A 10 ml injection of a sterile 1 mg/ml solution of 3 Evans blue was begun at exactly 65 minutes after starting H-PAH Injection.
Evans blue was injected during 30 seconds, and blood was aspirated into the injection syringe and re-injected three times. The 75 and 90 minute 52
plasma samples were used for the determination of Evans blue concentration. 3 3 To analyze for H-PAH and H-aPAH, blood filtrates were acidified with
concentrated H^PO^ (30 yl to 5 ml filtrate), tested with a pH dip-stick to
assure a pH of approximately 3.5, and injected into the HPLC as 100 yl
aliquots. A 10 mM NaH^PO^ buffer, pH 3.5, with 30% methanol was used as
the mobile phase at 2.5 ml/min. Eluate was collected during elution of
PAH and aPAH peaks, which were quite large from the PAH and aPAH spike
added to the blood immediately after collection. Under these conditions,
PAH eluted between 1:45 and 2:25 minutes; aPAH eluted between 2:30 and
3:15 minutes after injection. To the collected eluates, approximately 7
ml of Ready-Solv scintillation cocktail (EP type from Beckman) was added,
and samples were counted on a Beckman LS-8000 liquid scintillation
spectrophotometer with a window setting of 0-1000. The "pre-cold" and
"pre-hot" controls were used to determine background. DPM values were
calculated from CPM and the respective H number, and were used to cal
culate adjusted radioactivity values.
Compartmental analysis proceeded as described for the non-radioactive
single injection experiments. Adjusted radioactivity was calculated by
dividing the observed radioactivity by total dose radioactivity, and
multiplying by 100. Adjusted radioactivity was plotted on a log (base e) scale versus time, and are presented in Results and Discussion.
The apparent volume of PAH distribution was calculated from the
Aj^ and A^ values obtained from the compartmental analysis. The volume
of blood determined by Evans blue was compared to the PAH volume and
is discussed in the Results and Discussion. 53
Results and Discussion
For this discussion, "primary" denotes the compound that was injected, and "secondary" denotes the other compound. For example, in those experi ments in which PAH was injected, PAH is the primary compound, and aPAH is the secondary compound.
Experiments with non-radioactive PAH and aPAH
Both single-injection and continuous-infusion experiments were performed to attempt to describe the kinetics of PAH and aPAH excretion and/or interconversion. Following analysis and evaluation of data from these experiments, few conclusions could be made because of the large degree of variability in the adjusted concentration values and lack of reproducible results among experiments. The intent of this discussion is to present data from these experiments, consider possible sources of 3 variability and error, and describe how the H-PAH experiments were designed to avoid problems encountered during the non-radioactive experi ments.
Single injection The semilog plots of the adjusted concentration values for the single-injection experiments are in Appendix C. Figures
C.l - C.14 are those after PAH was injected; Figures C.15 - C.27 are those after aPAH was injected.
Slopes and intercepts (i.e., A^, kj^, A^, k^) estimated from data from single-injection experiments were of little value for quantitative information regarding the in vivo kinetics of PAH and aPAH. The graphs 54
in Appendix C reveal a large degree of variability in the results and a lack of reproducibility for both the primary and secondary compounds in both the PAH and aPAH experiments. This is especially evident in experi ments in which aPAH was injected.
A two-component exponential disappearance pattern is evident from the graphs; however, it is impossible to quantitatively calculate from the data. The concentrations of primary compounds for both PAH and aPAH single injections are reduced to approximately 20% of the initial concentration within the first few minutes after injection. A similar 14 pattern was reported for C-PAH by Oldendorf and Kitano (1972) in nephrectomized and non-nephrectomized rats. Their conclusion was that the rapid disappearance of PAH from plasma was due solely to equilibration with the extracellular space, and did not represent renal excretion. Van
Stekelenburg, et al. (1976) attributed the disappearance of hippuran to a three-component system, the most rapid component being explained by two peripheral compartments that were both freely connected to the plasma, but not to each other. Thus, hippuran rapidly equilibrated with these two pools within the first minute after injection.
The appearance of secondary compounds occurred shortly after the primary injection according to the graphs in Appendix C. A large degree of interference in the absorbance detection of PAH and aPAH was evident from blood filtrates collected during these experiments. Therefore, the secondary compound values are in question.
The sources of error and the large degree of variability encountered 55
in the single injection experiments will be considered in a general discussion of the non-radioactive experiments. The results of the single- 3 injection experiments were used to design the H-PAH experiments, and were not used for any quantitative information regarding either excretion or conversion rates.
Continuous infusion The continuous infusion approach for the study of in vivo kinetics of PAH and aPAH was an obvious choice for two major reasons. First, if PAH is to be used as a blood-flow marker for trans-liver balance studies, it probably would be administered as a continuous infusion to minimize minute-by-minute variations in blood flow.
Therefore, to validate PAH as a marker, it would be necessary to evaluate
PAH excretion rates and conversions to aPAH under a continuous-infusion situation. Second, if an acetylation reaction does occur in vivo In cattle, a constant, relatively high concentration of PAH would enable estimation of maximal acetylation of PAH. In other words, acetylation of
PAH would be more likely to be detected if the enzyme system was sat urated with PAH.
The blood concentration of infused compound in continuous infusions should reach a constant value when the amount of compound Infused equals the rate of removal of the compound from the blood. The GLM procedure of SAS (SAS Institute, 1979) was used to calculate least-squares linear regression estimates of the data points using samples collected after 20 minutes. Conclusive remarks about and equilibrium plateau are difficult to make from the predicted lines and observed points of the continuous infusion experiments graphed in Appendix D. The concentrations of PAH 56
and aPAH show linear patterns; however, because of the large degree of variation in the individual data points, quantitative information from these data cannot be calculated with confidence.
Other obvious problems with these data are inconsistencies observed for the aPAH continuous infusions. For four of the five experiments In which aPAH was infused, blood filtrates from the first few samples seemingly contained more PAH than aPAH (AC-1, Figures D.9-D.10; AC-2,
Figures D.11-D.12; AC-3, Figures D.13-D.14; and AC-4, Figures D.15-D.16).
It is unlikely that the infusion solution contained significant amounts of PAH, even though aPAH used in these experiments was synthesized from
PAH. Chromatograms of the infused solutions all had single peaks at the time aPAH standards elute; therefore, infusâtes were considered to be essentially pure.
Other problems believed to have contributed to the variable results are considered in a general discussion of the non-radioactive experiments, which follows.
General comments on the non-radioactive experiments Despite a lack of conclusive evidence from the single-injection and continuous infusion experiments, important information was obtained from the data.
Whether the variability resulted from the experimental treatments, animals, or analysis can only be speculated from these data; however, clues to the causes of the problems are evident.
First, one of the major problems with the design of the single- injection experiments was that the steers were not in steady-state with 57
respect to PAH or aPAH. Because these compounds are exogenous substances, their introduction into the animal's systemic circulation presents a
dynamic, constantly changing situation. As PAH (or aPAH) is removed from
circulating blood by any mechanism, the concentration in blood is continually decreasing. Such is not the case when a labelled endogenous substance is introduced as a single injection. Following a period of
mixing, the ratio of labelled molecules to endogenous molecules exiting the blood should be proportional to the amounts present. The declining concentration of PAH or aPAH in the blood potentially alters the rate of exit of these molecules from the blood. As discussed in the literature review of this dissertation, the renal transport of PAH changes as the concentration in the blood changes below the transport maximum; I.e., when transport follows first-order kinetics. Because it is generally accepted that the kidney is the sole site of PAH and aPAH removal, one
problem with single injections is an ever-changing transport rate of
PAH from blood to urine. In other words, it would be Impossible to reliably calculate excretion rates when the rates are not constant within
the animal.
Second, the concentrations of PAH and aPAH in blood samples were too low to quantitate accurately, especially during the continuous infusion experiments. When concentrations are low in blood filtrates, a large amount of baseline interference occurs from the high sensitivity settings of the absorbance detector of the HPLC and from unknown constituents in blood filtrates. This problem is probably the single most Important source of the variability and lack of conclusive evidence. 58
A primary cause of low blood concentration is merely that the doses used were too low for systemic administration. As an example, excretion of PAH will be considered for a 100 kg steer using a transport maximum value of 2.0 mg/(min x kg)reported for sheep by Rabinowitz, et al. (1971).
No other value for ruminants for the removal of PAH in vivo could be found in the literature. A 100 kg steer could excrete 200 mg of PAH/min , or 12.0 g/hr. For single injection experiments, 15, 30, and 45 mg were injected per lb BW; therefore, a 100 kg steer would have received 9.9 g at the 45 mg dose level. If the entire systemic blood supply passed through the kidneys with each circulation, the entire dose could be removed within a minimum of 50 minutes. This would be assuming that the maximum trans port of PAH would be occurring during the entire 50 minutes; in reality this would not be the case. When concentrations in the blood fell below that of the transport maximum of PAH, the transport of PAH into the urine would become proportional to the PAH concentration. Also, only about 25% of the cardiac output is circulated through the kidneys, thus prolonging the excretion of PAH from the blood. Thus, the entire 9.9 g dose in the
100 kg steer would take substantially longer than 50 minutes.
Gldendorf and Kitano (1972) provide evidence that PAH in rats is not confined to blood, rather it is rapidly diffused throughout the general extracellular space. In their report, ^H-mannitol and ^^C-PAH had virtually identical disappearances in nephrectomized and non- nephrectomized rats within the first minute of injection. Because mannitol is well-known as an extracellular space "marker", they concluded 59
that PAH also rapidly equilibrates with extracellular space. In non-
nephrectomized rats, PAH removal from blood was much more rapid than
mannitol after 1 minute, reflecting the high efficiency of PAH removal by
the kidneys. Nephrectomized rats displayed almost identical concentration
patterns for mannitol and PAH after the 1 minute equilibration period.
Thus, the 9.9 g dose in the previous example would be distributed in a space greater than the 8% BW estimate of blood mass; i.e., 8 liters for a 100 kg steer. Considering a 12% BW estimate for extracellular space, the 9.9 g dose would produce a concentration of roughly .825 mg/ml.
The dose levels for the single injection experiments were patterned after Mandel, et al. (1955), who used a dose rate of 4 mg/kg BW for single injections in dogs. When the present single injection experiments were designed and executed, it was felt, in error, that 15, 30, and 45 mg/lb would be more than adequate to provide the data needed to evaluate
the kinetics in vivo.
An important consideration to keep in mind when evaluating the present single injection and continuous infusion experiments is that systemic blood was studied rather than an isolated system such as the hepatic circulation. In their method for measuring hepatic and portal venous blood flow with PAH, Katz and Bergman (1969a) infused PAH into a mesenteric vein and sampled from the portal and hepatic veins. Thus, PAH was not diluted into the entire volume of the vascular system.
The difficulties in analyzing such low blood concentrations of PAH 3 and the problems with steady state were considered as the H-PAH 60
experiments were planned. The subsequent discussion explains the 3 rationale behind the design of experiments with H-PAH.
3 Experiments with H-PAH
Two significant problems observed from the experiments with non- 3 radioactive PAH and aPAH were kept in mind when the H-PAH experiments were designed. First, an animal must be in steady state with regard to
PAH if accurate estimates of the excretion or disappearance rates of PAH are to be obtained. Second, PAH and aPAH are difficult to quantitate in systemic blood due to their dilution into the entire vascular system and due to their rapid rates of removal from blood by the kidneys.
A continuous infusion of unlabelled PAH at 4.5 g/hr provided a constant source of PAH so that a steady state should have been achieved.
Thus, the disappearance rates measured after a single injection of 3 H-PAH during the continuous infusion of unlabeled PAH should reflect a constant removal rate from the total PAH pool. The concentration of PAH would be relatively constant, and the rate of removal would not be changing because concentrations would not be changing within the blood. 3 The use of H-PAH provided an alternative means to observe PAH disappearance from blood in that the disappearance of radioactive PAH rather than a decrease of PAH concentration could be monitored. The 61
experiments did not depend upon PAH and aPAH absorbance at 270 nm on the spectrophotometer used with the HPLC system for quantitation, and thereby avoided the interference encountered before. Additional unlabelled
PAH and aPAH added to blood samples immediately after collection produced large peaks during chromatography of blood filtrates; collection of 3 3 H-PAH and H-aPAH was easily accomplished.
A summary of steers, body weights, and experimental doses is given in Table 7. Body weights averaged 204 lbs. among the six animals; the range in weight was 145 to 240 lbs. Because total continuous infusion rates of unlabelled PAH were identical for all steers, relative rates
(mg/min «kg) changed with body weights. The mean relative rate of 3 unlabelled PAH infusion for experiments using H-PAH was 0.84 mg/min kg.
By comparison, the relative dose rate for continuous infusion experiments, described earlier, using only unlabelled PAH was 0.25 mg/min • kg ; the 3 dose rates for unlabelled PAH in the experiments with H-PAH were intentionally increased so that PAH steady state should be achieved in all steers.
The ^H-PAH dose rate had a range of 5.4 to 11.9 yCi/kg BW and averaged 8.29 yCi/kg BW. This dose level was designed so that adequate radioactive counts would be collected from blood filtrates for 3 H-aPAH following separation on the HPLC,
Observed adjusted radioactivity values and predicted lines for each experiment appear as semilog plots in Figures 6-11. The PAH lines were calculated from slopes and intercepts discussed in the following 62
Table 7. Steers, body weights, and dose rates used for experiments using ®H-PAH.
Body Unlabelled _ Steer Weight PAH dose^ H-PAH dose®
(lb.) (mg/min*kg) (mCi) (liCi/kg)
8446 240 0.69 0.78 7.14
8451 235 0.70 0.79 7.36
8452 214 0.77 1.14 11.90
8466 217 0.76 0.53 5.42
8483 145 1.14 0.41 6.17
8484 172 0.96 0.92 11.75
Mean 204 0.84 8.29 S.E. 15.3 0.07 1.15
^Administered as a continuous infusion of a 30 mg/ml PAH solution, pH 7.3, at a rate of 2.5 ml/min.
Adminstered as a single injection, in a total volume of 40 ml of 0.85% saline. Figure 6. Sendlogarithinic plot of observed adjusted radioactivity values and the predicted lines for PAH and aPAH following the injection of 0.78 mCi of ^H-PAH into steer #8446 while receiving a continuous infusion of unlabelled PAH at a rate of 4.5 g/hr. The equation describing the PAH disappearance is Adjusted Radioactivity = (1.42 x 10 '*)e + (3.07 x 10 ®)e
The equation describing aPAH appearance and disappearance is
Adjusted Radioactivity = (-2.52 x I0~®)e-o-37t + (2.52 x 10~5)e"°- 8%%6 PREDICTED VS OBSERVED PAH OBSERVED flPfiH
< E O = 5 - < -
0.00 2.00 6.00 8.00 10.00 12.00 TIME (MINUTES) (x 10*) Figure 7. Sendlogarithinic plot of observed adjusted radioactivity values and the predicted lines for PAH and aPAH following the injection of 0.79 mCi of ^H-PAH into steer #8451 while receiving a continuous infusion of unlabelled PAH at a rate of 4.5 g/hr. The equation describing the PAH disappearance is Adjusted Radioactivity = (1.38 x 10 '*)e + (6.64 x 10 ®)e The equation describing the aPAH appearance and disappearance is Adjusted Radioactivity = (-5.27 x 10 ®)e "•'*7^+ (5.27 x 10 ®)e B%51 PREDICTED VS OBSERVED PAH X r OBSERVED flPRH
>• H o = oE 5 gj * G\ og UJ = * &: • D - -s Q <
Y 0-4
-l— _j _j 1 , 1 0.00 2.00 y.oo 6.00 8.00 10.00 12.00 TIME (MINUTES) (X10^) Figure 8. Semllogarithmic plot of observed adjusted radioactivity values and the predicted lines for PAH and aPAH following the injection of 1.14 mCi of ^H-PAH into steer #8452 while receiving a continuous infusion of unlabelled PAH at a rate of 4.5 g/hr. The equation describing the PAH disappearance is Adjusted Radioactivity = (1.68 x 10 '*)e + (4.54 x 10 ®)e
The equation describing the aPAH appearance and disappearance is Adjusted Radioactivity = (-4.69 x 10 ®)e + (4.69 x 10 ®)e 8*52 PREDICTED VS OBSERVED PAH X OBSERVED fiPfiH •
as 00
1 r r 1 T \ 1 0.00 2.00 M.OO 6.00 10.00 12.00 TIME (MINUTES) 10^) Figure 9. SemlLogarithmic plot of observed adjusted radioactivity values and the predicted lines for PAH and aPAH following injection of 0.53 mCi of ^H-PAH into steer #8466 while receiving a continuous infusion of unlabelled PAH at a rate of 4.5 g/hr. The equation describing the PAH disappearance is
Adjusted Radioactivity = (2.29 x 10 '*)e + (4.50 x 10 ®)e
The equation describing the aPAH appearance and disappearance is
Adjusted Radioactivity = (-5.02 x 10 ®)e + (5.02 x 10 ®)e 8166 PREDICTED VS OBSERVED PAH OBSERVED flPfiH
>• -
6: <_
cc =
OE
?-
0.00 2.00 6.00 8-00 10.00 12.00 TIME (MINUTES) (xlO^) Figure 10. SemLiogarithinic plot of observed adjusted radioactivity values and the predicted lines for PAH and aPAH following injection of 0.41 mCi of ^H-PAH into steer #8483 while receiving a continuous Infusion of unlabelled PAH at a rate of 4.5 g/hr. The equation describing the PAH disappearance is
Adjusted Radioactivity = (3.53 x 10 '*)e + (8.93 x 10 ^)e
The equation describing the aPAH appearance and disappearance is
Adjusted Radioactivity = (-1.40 x 10 ®)e + (1.40 x 10 ®)e O'OZzt^ 8483 PREDICTED VS OBSERVED PAH X OBSERVED flPflH ^
X N3 •
I 1 M.00 12.00 TIME Figure 11. Eemilogarithmic plot of observed adjusted radioactivity values and the predicted lines for PAH and aPAH following injection of 0.92 mCi of ^H-PAH into steer #8484 while receiving a continuous infusion of unlabelled PAH at a rate of 4.5 g/hr. The equation describing the PAH disappearance is
Adjusted Radioactivity = (2.17 x 10 '*)e + (3.77 x 10 ®)e
The equation describing the aPAH appearance and disappearance is
Adjusted Radioactivity = (-4.35 x 10 ®)e + (4.35 x 10 ®)e 8%8% PREDICTED VS OBSERVED PAH OBSERVED flPRH
H"
O -
o—
D-
o
0.00 2.00 6.00 8.00 10.00 12.00 TIME (MINUTES) (x 10*) 75
paragraph. The predicted lines for aPAH curves are considered in a later 3 discussion. The use of two-component functions to describe H-PAH and 3 3 H-aPAH concentrations following H-PAH injection is justified because the semilog plots of data points are curvilinear. The predicted curves describe the observed values quite well for two experiments (#8452,
Figure 8; and #8483, Figure 10); predicted lines for the other experiments do not fit the observed data as well. Part of the difficulty in curve fitting of these data is caused by the low radioactivity encountered in 3 samples collected at later times, especially for H-aPAH. Many of the adjusted radioactivity values used to estimate the slope and intercept for the second component were 10 DPM or less above background; therefore, substantial errors may exist for these values.
Because the rapid component slope and intercept are calculated from residual values obtained after subtracting slow component values from observed values, an inaccurate estimation of the slow component would understandably result with an inaccurate estimate of the rapid component.
The estimated slopes and intercepts for the rapid and slow components 3 for H-PAH disappearance from blood appear in Table 8. These values were calculated by the graph analysis described in Materials and Methods.
The A^ values are the intercepts for the rapid component of the function that predicts the adjusted radioactivity for each time period;
Y = A e'^l^ + A e"^2t 1 2. •
The value is the functional disappearance rate constant and is equal 76
Table 8. Estimates of intercepts and slopes for the disappearance of ^H-PAH from blood®
Steer S ^2
8446 1.42 X 10-4 0.19 3.07 X lOT^ 0.051
8451 1.38 X 10-4 0.17 6.64 X loT^ 0.072
8452 1.68 X 10-4 0.21 4.54 X 10-5 0.053
8466 2.29 X 10-4 0.20 4.50 X 10-5 0.051
8483 3.53 X 10-4 0.16 8.93 X 10-5 0.029
8484 2.17 X 10-4 0.19 3.77 X 10-5 0.029
Mean 2.08 X 10-4 0.20 5.24 X 10-5 0.048
S.E. 0.33 X 10-4 0.007 0.89 X 10-5 0.007
^These estimates are used in the following function to describe the disappearance of %-PAH from blood:
Y = A^e'^l^ +
^The A and A values are the intercepts of the lines on the semilog plots describing the rapid and slow components, respectively. The units for A^ and A^ are 1/ml.
^The and values are the negative slopes of the lines on the semilog plots describing the rapid and slow components, respectively. The units for k^ and k^ are 1/min. 77
in value, but opposite in sign, to the slope of the straight line of the rapid component on the semilog plot. It can be interpreted to be the 3 fractional rate at which H-PAH disappears from blood by the rapid 3 component; i.e., an average of 20% of the existing H-PAH disappeared per minute. This corresponds to a half-time of 3.5 minutes, the time that 3 the H-PAH concentration was decreased in half by the rapid component. 3 The slow component for removal of H-PAH is evident in the semilog plots of the adjusted radioactivity vs time as the linear portion of the curves at times greater than 20 minutes. The intercepts and slopes averaged 5.24 x 10"^ and 0.048, respectively for the six experiments.
The fractional disappearance constant of 0.048/min corresponds to a half-time of 14.4 min. Thus, the second component appears to be approxi mately one-fourth as fast as the rapid component. A greater amount of variation is evident in the slow component values when compared to the rapid component values. As mentioned previously, many of the data points at later samples are based upon very low DPM values. The increased variation of the slow component slopes and intercepts probably is caused from the variation introduced by the low radioactivity.
Half-times for the removal of PAH have been reported to be 28 minutes in humans (Newman et al., 1949) and 40 and 33 minutes in dogs
(Blaufox et al., 1963; and Carlson & Kaneko, 1971; respectively).
These values were calculated from two to four data points and do not consider two components separately; therefore, it is difficult to compare these values to the 3.5 and 14.4 min half-times calculated from 78
the present data. However, the literature values for PAH removal half- times are significantly slower than those observed for either component of the present study. The discrepancies between the values for bovines and those from humans and dogs may be a difference in PAH-removal mechanisms in the different species, or differences in the experimental procedures. It is unlikely that experimental differences would cause such large discrepancies.
Figures 6-11 also contain the semilog plots of adjusted radioactivity 3 3 and predicted lines versus time for H-aPAH that is formed from H-PAH.
The rise and subsequent fall of the plot of these data points are typical of a two-component model (Shipley & Clark, 1972), in which the compound constituting the second component (aPAH) arises from the compound constituting the first component (PAH). With present data alone, it is impossible to assess the reversibility of the PAH to aPAH reaction; experiments with labelled aPAH would be required. But, the graphs clearly indicate a rapid acetylation of PAH. The slopes and intercepts for the appearance and disappearance components for the aPAH adjusted radio activity predicted curve appear in Table 9. The A^ and A^ values for this curve are of equal value, except that A^ is of opposite sign to A^.
The equation for the aPAH curve remains the same as for the disappearance of PAH, except that the A^ and A^ values are equal and A^ is of opposite sign to Ag. The average intercept for aPAH (5.97 x 10 ^) is approximately one-tenth that of the slow component for PAH (5.24 x 10 ^). It is 79
Table 9. Intercepts and slopes for the line predicting the appearance and disappearance of aPAH&
Steer Intercept^ kl' ^2
8446 2.52 X 10"^ 0.37 0.032
8451 5.27 X lOT* 0.47 0.033
8452 4.69 X 10"^ 0.31 0.072
8466 5.02 X 10"^ 0.23 0.031
8483 1.40 X 10"^ 0.26 0.022
8484 4.35 X 10"^ 0.43 0.021
Mean 5.97 X 10"* 0.35 0.035
S.E. 1.65 X 10"* 0.039 0.008
^The equation is Y = A^e + A^e ^2^,
^The intercepts, A^^ and A^ are equivalent except that A^ is negative and Ag is positive. A^^ and A^ have units of 1/ml.
^The k]^ and k2 values are equal and opposite in sign to the slopes of lines of the appearance and disappearance, respectively. Both values have units of l/min. difficult to assign significance to this finding without investigations using aPAH as the primary compound. However, values for the aPAH curve averaged ten times more than values. This implies aPAH appearance occurs at a rate ten times that as for aPAH disappearance from blood.
As before, it is hard to fully interpret these rates without data concerning the disappearance of aPAH from blood after aPAH injection.
Comparison of the slow component for PAH and the disappearance component for aPAH indicates that PAH was removed at a faster rate than was aPAH (half-times were 14.7 for the PAH slow component and 19.8 for the second component for aPAH). It must be remembered when interpreting these values that PAH is converted to aPAH, and the disappearance rate reflects total PAH removal. Therefore, the fractional disappearance constants for PAH is a combination of removal by conversion to aPAH and removal by excretion. The values for aPAH and PAH removal rates may be closer together if aPAH synthesis from PAH could be accounted for.
Although PAH is obviously acetylated to aPAH in steers, there does not appear to be a major conversion of PAH to aPAH. The plots presented in Figures 6-11 are graphed on a semilog scale. Thus, the PAH and aPAH values appear to be close together when graphed, when actual values for aPAH are 1/100 or 1/10 that of PAH. As before, many of the later samples contained relatively low radioactivity and variability in these samples among experiments is more evident. 3 Analysis of urinary excretion of H indicated an average recovery of 88.5% of the amount injected from all experiments, as indicated in 81
Table 10. These results must be viewed with some caution, however, 3 because they are based on a relatively crude measurement of H-excretion.
Several potential sources of error are possible. First, a graduated cylinder was used for measurement, and urine volumes were recorded to the nearest ml. Second, some steers (e.g., #8446 and #8452) positioned themselves so some urine missed the collection pans and was lost on the floor. Third, the collection pan had a screen to prevent particles of fecal material from clogging the funnel. Cross-contamination of successive urinations could easily occur from residual radioactivity in the collection system. Despite thorough water washings between experiments, cross-contamination surely happened. Radioactive counts of approximately 2,000 dpm per ml of urine were obtained in pre- injection urine samples in four experiments (#8446, #8451, #8466, and 3 #8484). Fourth, the time lag between removal of H-PAH from blood, storage in urine, and micturation was quite varied. One steer (#8452) did not urinate for over 4 hours after collection of the last blood sample.
Probably much of the uncollected radioactivity was excreted after the steer was removed from the crate.
The PAH volume, relative PAH volume, blood volume, relative blood volume, and PAH volume : blood volume ratios are listed in Table 11. The
PAH volume has been defined as the apparent space in which PAH is distributed and was calculated from the sum of the A^ and A^ values listed in Table 8. The sum of the intercepts is the theoretical value of the adjusted radioactivity if Instantaneous mixing had occurred at 82
3 Table 10. Radioactivity recovered in urine after H-PAH injections
Steer Dose Excreted Recovery
(dptn) (dpm) (%)
8446 1.73 X 10^ 1.34 X 10® 77.5
8451 1.75 X 10® 1.62 X 10® 92.6
8452 2.52 X 10® 1.97 X 10® 78.2
8466 1.19 X 10® 1.38 X 10® 116.1
8483 9.02 X 10® 8.82 X 10® 97.8
8484 2.04 X 10® 1.41 X 10® 69.1 Mean 88.5 S.E. 7.6 83
Table 11. Volumes of distribution of PAH and blood volume from Evans blue
PAH volume; PAHS Relative Blood^ Relative blood volume Steer volume PAH volume volume blood volume ratio
(1) (1/kg) (1) (1/kg)
8446 22.4 20.5 11.4 10.5 1.95
8451 18.8 17.6 13.2 12.4 1.42
8452 18.1 18.6 12.6 12.9 1.44
8466 14.1 14.3 11.7 11.8 1.21
8483 8.7 13.2 7.6 11.6 1.14
8484 15.2 19.4 10.5 13.4 1.44
Mean 14.5 17.3 12.1 1.43
^PAH volume was calculated from the A^ and A^ values from Table 8.
^Blood volume was determined according to the Evans blue procedure described in Appendix B. 84
time 0. The PAH volume averaged 14.5 1 for the six experiments. A more useful figure for comparing PAH volumes is the relative PAH volume, or liters of PAH volume divided by kg BW, which averaged 17% of the BW.
Relative blood volume, the blood volume measured by Evans blue dilution divided by kg BW, averaged 12.1. This blood volume appears relatively high compared to the accepted estimate of 8% of the body and seems to more closely approximate the 12% BW estimate for extracellular space.
Evans blue is considered to be confined to blood after injection; its use as an indicator for blood volume is well-established. Blood volume would be over-estimated when the dye appears to be diluted more than it would be by the blood alone. Several explanations can be offered for this apparent over-dilution. First, plasma blanks used to zero the spectro photometer for Evans blue quantitation may have been slightly hemolyzed.
Hemolyzed plasma would zero the spectrophotometer at higher absorbances than would non-hemolyzed plasma. Evans blue would then appear less concentrated than it actually was; blood dilution would be overestimated.
Second, problems with standard curve absorbances possibly could cause underestimation of Evans blue concentrations. No notable differences among standard curves were noted, although no in-depth analysis was conducted to analyze standard-curve differences.
The ratio of PAH volume to Evans blue blood volume ranged from
1.14 for steer #8483 to 1.95 for steer #8446 and averaged 1.43 among the six steers. This indicates that PAH is distributed in a space larger than blood volume. Even though the volume of blood is overestimated. 85
PAU is distributed in a larger space. Caution must prevail, however,
because the apparent distribution of PAH not only reflects the physical
space, i.e., mixing, but also the chemical space, i.e., transformation
to other compounds. In other words, the volume of distribution is
calculated from the intercepts of the adjusted radioactive curve and dose,
and, therefore, is a composite of all factors which dilute PAH. To
assign a physical space to this distribution would be unfounded from
these data alone, although dilution into a space greater than that of
blood is suggestive that PAH is not confined to blood. A similar distri
bution was reported in rats by Oldendorf and Kitano (1972). Mandel, et al. (1955) reported a PAH distribution in rats representing 25.6% of the body weight, and concluded that PAH may be penetrating some intracellular space. The 17% BW distribution observed in the current studies would not indicate such a distribution.
Conclusions
The results reported in Part Two of this dissertation have provided
information that allows the following conclusions to be reached. First,
acetylation of PAH does occur in vivo in cattle but the actual
magnitude cannot be calculated for these steers. Actual rates of
acetylation cannot be determined from these data alone, although aPAH
appearance in the blood was evident within 5 minutes of injecting PAH,
and reached maximum amounts between 15 and 25 minutes. The appearance
of aPAH occurred at a fractional appearance rate of .3465, indicating
a Ti time of appearance of 2.0 minutes. Second, PAH does not appear to
be confined to blood after injection into systemic blood. The distri 86
bution corresponds more closely to the extracellular space. Third, approximately 90% of the injected radioactivity was recovered from 3 urine within 5 hours after injection of H-PAH.
The present results do not confirm or invalidate the use of PAH as a blood-flow marker. It is evident that aPAH must be considered or minor errors would result In flow calculations. Further experimentation is necessary to evaluate rates of acetylation. But, the acetylation of PAH does not appear to be a major means by which PAH is removed from blood.
Also, investigations with PAH and aPAH in the portal system need to be conducted before their use in portal-flow determinations can be validated. Not being able to obtain specific activity values for PAH 3 and aPAH after the H-PAH injection greatly reduced the amount of 3 information that could be derived from experiments with H-PAH. 87
POSTSCRIPT
The use of p-amlnohippurate as a marker in steers remains an
attractive means to determine blood-flow for trans-hepatic balance
studies. The HPLC procedure developed for PAH and aPAH provides an
easy, reliable analytical procedure that can be routinely performed.
Deproteinization of blood samples with H^CIO^ and KOH is the only
preparation required prior to analysis and detection by absorbance at
270 nm does not require derivitizing agents.
PAH is rapidly coverted to aPAH in vivo; however, this does not appear a major route of PAH removal from the blood. Because the
primary site of PAH acetylation is believed to be the kidney, aPAH that is formed would have negligible effects on blood flow calculations for the hepatic circulation.
Because the liver has very little functional extracellular space, the fact that PAH presumably is distributed in extracellular space would not in itself alter flow calculations. Flow calculations would not be invalidated even if significant extracellular distribution occurs; an equilibrium between the vascular and extravascular space would exist in continuous infusion conditions after the distribution is established. 88
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ACKNOWLEDGEMENTS
My gratitude goes to Dr. Jerry W. Young for his countless hours of patience and guidance while I struggled through the last three
years of graduate school. His efforts to make me a responsible scientist hopefully will be fruitful. Dr. Donald Beitz has been and will continue to be an inspiration to me; I hope my career is as successful as his has been. I'm proud to call him "friend" as well as "teacher". The surgical skills and guidance provided by
Dr. Dare McGilliard will benefit me for a long time. To Drs. Jeanine
Carithers and James Redmond, thank you for consenting to join my committee for evaluating the dissertation and conducting the final examination. In addition, the teachings and inspiration of the late
Dr. David R. Griffith will be a guide to my career; he is sorely missed.
I gratefully acknowledge Eli Lilly and Co. of Indianapolis,
Indiana and Grant #901-15-160 funded under special grants program of
SEA-CR for financial assistance.
I thank my fellow graduate students for many stimulating discussions and helpful criticism, I wish them the best of luck in graduate school and their careers.
Finally to friends and family; my brothers, sisters, and especially my parents, thank you for your support and confidence.
Most of all, thanks to my wife, Pam. She has an endless supply of love and patience. 99
APPENDIX A. PEAK-HEIGHT RATIO PROCEDURE FOR QUANTITATION
PAH AND aPAH USING FAB AS AN INTERNAL STAITOARD
The following steps were performed for quantitation of PAH and aPAH samples containing unknown concentrations:
1. The detector response constancy was determined by measuring peak
heights of all three compounds from standard solutions. The PAB
internal standard peak height is divided by the peak heights of
PAH and aPAH to produce the PAB:PAH (P-Std Ratio) and PAB: aPAH
(A-Std Ratio) standard ratios.
2. A known amount of PAB is added to the blood sample immediately
after collection. The volume and concentration of the PAB
solution must be known, but numerous combinations of volume and
concentration are acceptable.
3. The blood samples are analyzed on the HPLC and the peak heights
for all three compounds are measured. The PAH:PAB (P-Unk Ratio)
and aPAH:PAB (A-Unk Ratio) peak-height ratios are calculated by
division of the appropriate peak heights by the PAB peak height.
4. The total unknown amount contained in the entire sample volume
is calculated according to the following equations:
pmoles PAH = (P-Unk Ratio) x (P-Std Ratio) x total ymoles PAB added. ymoles aPAH = (A-Unk Ratio) x (A-Std Ratio) x total ymoles PAB added. 100
Comments
1. Note that the "Std Ratios" are in a reciprocal relationship
to the "Unk Ratios"; i.e., the "Std Ratios" are PAB divided
by PAH or aPAH, whereas the "Unk Ratios" are PAH or aPAH divided
by PAB.
2. The amount of PAH or aPAH obtained from the equations is the
total amount, inymoles, contained in the total amount of
sample collected. 101
APPENDIX B. EVANS BLUE PROCEDURE
A. Standard Curve
1. Preparation of stock dye solution. a. Prepare 0.1% solution of Evans Blue in H„0 (i.e., 1.0 g/ liter). b. Filter with JENA or Pyrex sintered glass filter. The residual dye left on the filter is insignificant, and no attempt should be made to recover it. c. Place 11 ml in neutral glass ampules and seal. d. Autoclave at 121° for 20 minutes.
2. Calibration of the spectrophotometer. a. Dilute the stock solution ten times (i.e., 1:9) with 0.85% NaCl. This is the working solution. b. In duplicate, transfer 1, 2, 3, 4, 5, and 6 ml of the working solution to clean test tubes and dilute to 10 ml with 0.85% NaCl. c. Transfer 300 vil of each solution of dye to another tube and add 2.7 ml of plasma. d. Prepare the blank by mixing 300 yl of 0.85% NaCl and 2.7 ml plasma. e. Using 635 nm, zero the absorbance on the spectrophotometer with the blank. f. Read each dilution of dye at 635 nm and record the absorbance. g. Prepare a plot of the absorbance versus the dye concentration. The concentrations of the standard curve are 0.1, 0.2, 0.3, 0.4, 0.5, and 0.6 mg% (mg/dl).
B. Determination of total blood and plasma volume.
1. General Considerations. a. Use only dry syringes and needles. Avoid hemolysis, because hemoglobin interferes with the absorbance of Evans Blue. b. No dye must be lost in tissues; i.e., all dye must be injected into the bloodstream. c. Accurate timing is necessary; therefore, a stop-watch should be used for dye injection and blood sampling. d. It takes 4 to 6 minutes for complete mixing of the dye with the blood. The blood should not be sampled for at least 4 minutes after the injection of the dye.
2. Procedure a. Withdraw about 6 ml of blood from a vein. Fill 2 hematocrit tubes with blood and fill one end with sealing putty, then centrifuge. Place the remaining blood in tubes containing 1 drop of concentrated heparin. Make sure the resulting plasma is absolutely free from hemolyzed blood. This plasma is the blank. 102
b. Inject an exact amount of the dye solution (usually 10 ml). 1) the injection should take 30 seconds. il) draw blood back into the syringe at least 3 times and reinject the blood to wash out the residual dye in the syringe and needle. c. Exactly 10 minutes after the beginning of the injection, withdraw 6 ml of blood. Repeat #2a for this sample. d. Use the blank to zero the spectrophotometer at 635 nm. Read the sample and record the absorbance
3. Calculations a. Determine the mg % of dye in the plasma from the table constructed from the standard curve. b. A correction is necessary to account for the amount of dye absorbed by the tissues during the period from the injection of dye to the sampling of blood: Add 1.0% of the mg% of the dye in the blood for the 10 minute period. (Add 0.1% for each additional minute if the blood is with drawn at different times after injection.) c. Plasma volume;
Plasma volume (ml) - d'e'În plasma
d. Total blood volume:
Blood volume (ml) = [^(z'plcked^cells)
e. Calculate plasma and blood volumes on a kg basis by dividing values by kg B.W. 103
APPENDIX C.
PLOTS FOR SINGLE-INJECTION EXPERIMENTS
USING UNLABELLED PAH AND aPAH Figure C.l. Semilogarithinic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P15-3, performed on animal #8232 on September 10, 1980. The animal weighed 252 lbs. and received 3600 mg of PAH. WL. PIS 3 PAH CURVE % flPflH CURVE 'b Ul.
I.. "
V X • Œ rU tf o Ul
O % O tn- 4/ O • t o % 4> UJ (OJ m 5
O
o
1 \ 1 n 1 1 1 0.00 8.00 16.00 24.00 32.00 yo.oo y8.00 56.oo TIME (MINUTES) Figure C.2. Seoilogarithmic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P15-4, performed on animal #8241 on September 10, 1980. The animal weighed 234 lbs. and received 3400 mg of PAH. PIS % PflH CURVE in_ ' APAH CURVE
0-^ m-
I«H * * ° * z oô: O - 5- S H ^ * 4> X
Z ? • O 2 Q «H lU I
2 •" ta. Û 4/
Yo
1 1 1 1 1 0.00 2.00 y.00 6.00 8.00 10.00 12.00 TIME (MINUTES) ) Figure C.3. Senrilogarithmlc plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P15-5, performed on animal #8193 on September 15, 1980. The animal weighed 290 lbs. and received 5400 mg of PAH. PIS 5 PAH CURVE X APAH CURVE f-. 10- X L- X o X X z Id: O -
5"^ CE - X t
S"
D - 3- •
?
I I 1 1 1 1 1 ! 0.00 y.OO 8.00 12.00 16.00 20.00 24.00 28.00 TIME (MINUTES) ' Figure C.4. Sendlogarithinic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P15-6, performed on animal #8263 on November 6, 1980. The animal weighed 340 lbs. and received 5100 mg of PAH. PIS 6 PAH CURVE * flPflH CURVE tb
N
Z o ~
EU * lU g w *
8H ^
S "1 If f- V) m Q <
Y_ o
1 1 1 1 1 1 1— 0.00 U.OO 8.on 12.00 16.00 20.00 24.00 28.00 TIME (MINUTES) (xlO^ 1 Figure C.5. Setdlogarlthmlc plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P30-1, performed on animal #8193 on September 11, 1980. The animal weighed 290 lbs. and received 8800 rag of PAH. co P30 1 PAH CURVE flPflH CURVE in-
II OC X I- s (D- O - oz o^
Q * ÎÏCO, (/> 3 -9 Q ~ < I
f 1 1 1 1 1 0.00 4.00 8.00 12.00 16.00 20.00 2H.0a TIME (MINUTES) Figure C.6. Seudlogarithmic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P30-2, performed on animal #8241 on September 11, 1980. The animal weighed 234 lbs. and received 7000 mg of PAH. P30 2 PflH CURVE X "^3 APAH CURVE
C? ^ * Sr * X X * Zo tcD-
S- cc -} 5 J lU Ln o 2 •4/ O X ^ lU -
D -1 3«J <
o ^
1 1 1— _j J J 1 0.00 2.00 y.QO 6.00 8.00 10.00 12.00 14.00 TIME (MINUTES) J Figure C.7. Semilogarithmic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P30-3, performed on animal it8232 on September 11, 1980. The animal weighed 252 lbs. and received 7600 mg of PAH. P30 3 PAH CURVE in_ APAH CURVE
OQ- O - 5ud œ goj iU
X
< ? o
1 1 1 _j 1 1 0.00 4.00 8.00 12.00 16.00 20.00 2U.00 TIME (MINUTES) (xlO^ ) Figure C.8. Senrilogarithroic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P30-4, performed on animal #8232 on September 15, 1980. The animal weighed 266 lbs. and received 7600 mg of PAH. P30 4 PflH CURVE X ta. APAH CURVE X
X X z «c X o -
CC -1 X UJ ^ ^ ^ VO 4/ • Oil • Q°^ UJ - H la. (0 3 4y gtnj < f o
1 1 1 -| ! 1 1 .00 2.00 y.00 6.00 8.00 10.00 12.00 U.OO TIME (MINUTES) î*10' ) Figure C.9. SenrLlogarithmic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P30-4, performed on animal #8241 on November 6, 1980. The animal weighed 308 lbs. and received 9200 mg of PAH. P30 5 PflH CURVE X i APAH CURVE
i 4> 5 fiC H Z: ro lU o Z' o 4/ o • Q I U1 I— tn 3 -) O <
fj o
1 1 1 -i 1 1 1 0.00 y.OO 8.00 12.00 16.00 20.00 2%.00 28.00 TIME (MINUTES) (xlQi ) Figure C.IO, Sendlpgarithmic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P45-1, performed on animal #8193 on July 21, 1980. The animal weighed 265 lbs. and received 11940 mg of PAH. pits 1 PflH CURVE IA_ APAH CURVE if * X z tdq * O
5 J ^ * Ul 4xV * gs
X 4> si 4/
3^
f D
1 1 1 1 1 1 0.00 y.OO 8.00 12.00 16.00 20.00 24.00 TIME (MINUTES) (xlO* 1 Figure C.ll. Sendlogarithmic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P45-2, performed on animal #8229 on July 24, 1980. The animal weighed 175 lbs. and received 8000 mg of PAH. pits 2 d PAH CURVE X Ul_ flPflH CURVE •
X X X zbô: O - X 4% X K 4x • ro Ul lU • 4>
4/
Ul 4> H- Ul. • (0 3 3^ < Y" o
1 1 1 1 1 I I 0.00 «1.00 8.00 12.00 16.00 20.00 2U.00 28.00 TIME (MINUTES) xlQi 3 Figure C.12. Senrilogarithmlc plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P45-3, performed on animal #8193 on July 24, 1980. The animal weighed 276 lbs. and received 21420 mg of PAH. OCUi P45 3 PflH CURVE la. APAH CURVE
S^ l?1 * 2 *
g -
2^ ^4 $ Ul A, Seëf $ $ 04
S°3 4, • 5-^ 4/ 3 i 3"J < ?" o
J 1 1 T 1 1 r- 0.00 y.00 8.00 12.00 16.00 20.00 24.00 28. TIME (MINUTES) (xlQi I Figure C.13. Sendlogarithinic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P45-4, performed on animal #8241 on July 24, 1980. The animal weighed 194 lbs. and received 8800 mg of PAH. P«45 II PflH CURVE in» APAH CURVE
m k X X «-1 Z m: * o - X s- Œ ^ * X UJ • 4/ §s' X Ox Q«g UJ - •
3 gc^ < V" D
1 1 1 1 1 1 0.00 H.OO 8.00 12.00 16.00 20.00 2U.0a TIME (MINUTES) (*10^ ) Figure C.14. SemLlogarithmic plot of observed adjusted concentration values for PAH (asterisks) and aPAH (diamonds) versus time for experiment P45-5, performed on animal #8193 on July 28, 1980. The animal weighed 280 lbs. and received 12600 mg of PAH. PilS 5 PAH CURVE X la. flPflH CURVE
It- X X zed: g - X H Ul_ cc -|
UJ * UJ §B ox h X ^ X UJ - tr (b 5"^ X 3 I 4/ < f" o
1 1 1 1 1 1 1 0.00 4.00 8.00 12.00 16.00 20.00 24.00 28.00 TIME (MINUTES) 1*10* ) Figure C.15. Sendlogarithtnic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A15-1, performed on animal #8193 on July 20, 1980. The animal weighed 265 lbs. and received 4000 mg of aPAH. R15 1 PAH CURVE in_ APAH CURVE
L-o
o"^ %
OC -
4> W UJ _ w Ë9-m ^ * OS 4> * Ox * X ^ 0«N UJ
n
Î o <-4X 1 1 1 1 1 1 \— 0.00 4.00 8.00 12.00 16.00 20.00 2U.00 28.00 TIME (MINUTES) IxlO^ J Figure C.16 Semiltjgarithmlc plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A15-2, performed on animal #8232 on September 30, 1980. The animal weighed 270 lbs. and received 4000 mg of aPAH. co A15 2 PAH CURVE X APAH CURVE in.
p w UJos- Ln oË-_ - Om- • Û UJ
O -. * < T o
1 1 1 "T 1 1 1 0.00 2.00 y.OO 6.00 8.00 10.00 12.00 m.oo TIME (MINUTES) txio' J Figure C.17. Semilogarithniic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment Â15-3, performed on animal #8232 on September 26, 1980. The animal weighed 270 lbs. and received 4000 mg of aPAH. 4> CO- HIS 3 PAH CURVE X <%> APAH CURVE 'Si-
Z . O
IIZ^
O lU *
3 -3 o < m Î o
1 1 1 "I 1 1 1 0.00 2.00 y.OO 6.00 8.00 10.00 12.00 m.oo TIME (MINUTES) (xlO^ 1 Figure C.18. Semiiogarithmic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A15-4, performed on animal #8193 on September 3, 1980. The animal weighed 290 lbs. and received 4400 mg of aPAH. filS >4 PAH CURVE
•
K
X
* *
1 1 1 1 1 2.00 U.DO S.OO 8.00 10.00 12.00 TIME (MINUTES) 1*10^ J Figure C.19. Senrilogarithmic plot of observed percent adjusted concentration values for aPAH (diamonds and PAH (asterisks) versus time for experiment A30-1, performed on animal #8193 on September 19, 1980. The animal weighed 290 lbs. and received 8800 mg of aPAH. cnj A3D 1 PAH CURVE X APAH CURVE •
* Is 0 * ^ * ^ Z to. O - 5- X oc - *
o"5 X UJ
^cru
T D
f 1 1 _j J ; ; 0.00 2.00 y.QO 6.00 8.00 10.00 12.00 lU.OO TIME (MINUTES) [xlO^ J Figure C.20. Semilpgarithmic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A30-2, performed on animal #8232 on September 19, 1980. The animal weighed 252 lbs. and received 7600 mg of aPAH. A30 2 PRH CURVE * flPflH CURVE
it
X
• •
W
*
_1 1 ; 1 1 1 1 0.00 8.00 16.00 2U.00 32.00 40.00 ye.00 56.oo TIME (MINUTES) Figure C.21. Serailogarithinic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A30-3, performed on animal #8193 on October 1, 1980. The animal weighed 290 lbs. and received 9800 mg of aPAH. A30 3 PAH CURVE X flPflH CURVE
|t~ » 0 X % _
ZocZ 2 I *
CC - •
4> s"^ * * * Ui is' * "" w si 5"^ D -
<
1 <&
1 1 1— 1 1 1 1 0.00 2.00 y.OO 6.00 8.00 10.00 12.00 U.OO TIME (MINUTES) txlO^ Î Figure C.22. Senrilogarithmic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A30-4, performed on animal #8232 on October 1, 1980. The animal weighed 270 lbs. and received 8200 mg of aPAH. A3D «4 PflH CURVE APAH CURVE
^ -1^ C
IJ \ »
O O
Q lU 5 pj * (b * ^ •
f o 1 1 1 "T 1 1 0.00 y.oo 8.00 iz.aa 16.00 20.00 2%.00 TIME (MINUTES) (xlOi ] Figure C.23. SemLlogarithmic plot of observed percent adjusted concentration values for aPAH (diamonds and PAH (asterisks) versus time for experiment A30-5, performed on animal #8263 on November 7, 1980. The animal weighed 340 lbs. and received 10200 mg of aPAH. - A30 5 PAH CURVE uv APAH CURVE 1 I 1 1 -| 1 1 0.00 4.00 8.00 12.00 16.00 20.00 24.00 TIME (MINUTES) (xlQi } Figure C.24. Seœilogarithmic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A45-1, performed on animal #A45-1 on Ju]y 20, 1980. The animal weighed 210 lbs. and received 9450 mg of aPAH. — ms 1 PRH CURVE X in- APAH CURVE 4> fO- E \ ?" o €-* 4> X r-i_ z (dl o — 1- < in. HDC — Z Ln lU I—• o — * X 1" oz u * o X 00- « lUa H in. 3O) -3Q Pi. < o «-• X "T 1 ! I 0 y.oo 8.00 12.00 16.00 20.00 24.00 28.00 TIME (MINUTES) (xlQi 1 Figure C.25. Semilogarithinic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A45-2, performed on animal #8241 on October 3, 1980. The animal weighed 260 lbs. and received 11800 mg of aPAH. Alts 2 4!> PAH CURVE X APAH CURVE • Z "" X g _i Q lU W z 3 - 4? -9 o < o 1 1 1 1 1 1 1 0.00 y.OO 8.00 12.00 16.00 20.00 2%.00 28.00 TIME (MINUTES) UIO^ ) Figure C.26. SendLogarithmic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A45-3, performed on animal #8232 on October 3, 1980. The animal weighed 270 lbs. and received 12200 mg of aPAH. ms 3 PAH CURVE X APAH CURVE » o 5- oc - Ln Ln if • • * 1 i 1 _j ; 1 1 0.00 y.oo 8.on 12.00 16.00 20.00 24.00 28.00 TIME (MINUTES) (xlQi I Figure C.27. Semilogarithtnic plot of observed percent adjusted concentration values for aPAH (diamonds) and PAH (asterisks) versus time for experiment A45-4, performed on animal #8241 on November 7, 1980. The animal weighed 308 lbs. and received 13860 mg of aPAH. ms It PAH CURVE Ul_ APAH CURVE Pl. it 4> <î> ZttC o - 4> gwd oc -I U1 §B (b lU - K-in. (0 3 3^ < o i 1 1 1 1 1 r- 0.00 8.00 16.00 2%.00 32.00 yo.ao wa.oo 56. TIME (MINUTES) 158 APPENDIX D. PLOTS FOR CONTINUOUS INFUSION EXPERIMENTS USING UNLABELLED PAH AND aPAH Figure D.l. Plot of observed values and the predicted line for PAH concentration vs time for experiment PC-1, performed on steer #8263 on November 3, 1980. The animal weighed 340 lbs. and received a 775 mg primer dose of PAH and a continuous infusion of 38.7 mg PAH/min. The equation of the predicted line is PAH concentration = (9.3 x 10 x time + 0.134. o 3» PC 1 OBSEftVCD PRM vS' PREDICTED LINE X Ë8 m * w X UJ m D * M = n % X O a» d" I I r 1 r 1 1 0.00 y.00 8.00 ^if.oq 16.00 20.00 2y.00 28.00 TIME (MINUTES) (xio* J Figure D.2. Plot of observed values and the predicted line for aPAH concentration vs time for experiment PC-1. After a 775 mg primer dose of PAH, 38.7 mg of PAH/min was infused. The equation of the predicted line is aPAH concentration = (6.3 x 10 x time + 5.7 x 10 OBSERVED flPRH PREDICTED LINE «D O 0.00 8.00 12-00 16.00 20.00 28.00 TIME (MINUTES) (xio' ) Figure D.3. Plot of observed values and the predicted line for PAH concentration vs time for experiment PC-2, performed on steer #8263 on November 8, 1980. The animal weighed 340 lbs. and received a 775 mg primer dose of PAH and a continuous infusion of 38.7 mg PAH/min. The equation of the predicted line is PAH concentration = (-8.7 x 10 x time + 0.133. Q a» PC 2 OBSERVED PRH rJ" PREDICTED LINE zS ÎS-- w _J o Is * K * *- n X o I I 1 1 1 1 1 0.00 y.OO 8.00 12.00 16.00 20.00 2y.00 28.00 TIME (MINUTES) (xio^ J Figure D.4. Plot of observed values and the predicted line for aPAH concentration vs time for experiment PC-2. After a 775 mg primer dose of PAH, 38.7 mg of PAH/min was infused. The equation of the predicted line is -6 aPAH concentraton = (-1.1 x 10 ) x time + 0.017. PC 2 OBSERVED fiPRH + PREDICTED LINE Çg _l D o\ o\ 2: a- n o o Q" 0.00 4.00 (MINUTES)®'"" ^°(xîo» i Figure D.5. Plot of observed values and the predicted line for PAH concentration vs time for experiment PC-3, performed on steer #8241 on November 11, 1980. The animal weighed 310 lbs. and received a 700 mg primer dose of PAH and a continuous infusion of 35.3 mg PAH/min. The equation of the predicted line is _5 PAH concentration = (-3.3 x 10 ) x time + 0.073. PC 3 OBSERVED PRH % PREDICTED LINE UJ =tg X X n * X S- o Q Q 1 1 1 1 1 1 ; 0.00 y.OO 8.00 12.00 16.00 20.00 24.00 28.00 TIME (MINUTES) (xio' J Figure D.6. Plot of observed values and the predicted line for aPAH concentration vs time for experiment PC-3. After a 701 mg primer dose, 35.3 mg of PAH/min was infused. The equation of the predicted line is aPAH concentration = (-2.02 x 10 x time + 0.006. PC 3 OBSERVED fiPftH PREDICTED LINE O n 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) Cxio' i Figure D.7. Plot of observed values and the predicted line for PAH concentration vs time for experiment PC-4, performed on steer #8263 on December 4, 1980. The animal weighed 355 lbs. and received a 805 mg primer dose of PAH and a continuous infusion of 40.3 mg PAH/min. The equation of the predicted line is PAH concentration = (3.2 x 10 x time + 0.058. a PC It 3» OBSERVED PAH PREDICTED LINE S5-' LU _J O ag n Sg 5°^ * X "T 1 1 -| 1 1 0-00 y.oo 8.00 12.00 16.00 20.00 24.00 28.00 TIME (MINUTES) (xlQi ; Figure D.8. Plot of observed values and the predicted line for aPAH concentration vs time for experiment PC-4. After a 805 mg primer dose of PAH, 40.3 mg of PAH/min was infused. The equation of the predicted line is aPAH concentration = (-5.72 x 10 x time + 1.5 x 10 o PC 4 a OBSERVED flPflH PREDICTED LINE Çg E3 E •se s° o a H 4 "I—4 rH 4 T + -j H—I 4 f—H i -T" 0.00 y.oo 12.00 16.00 20.00 2(1.00 28.00 ®*nME (MINUTES) txio' Figure D.9. Plot of observed values and the predicted line for aPAH concentration vs time for experiment AC-1, performed on steer #8232 on November 14, 1980. The animal weighed 325 lbs. and received a 750 mg primer dose of aPAH and a continuous infusion of 36.9 mg aPAH/min. The equation of the predicted line is aPAH concentration = (1.51 x 10 x time - (8.9 x 10 ^). AC 1 OBSERVED RPflH PREDICTED LINE + % _J o 2: "^9 n + a a + d i" ' I I I I 1 1 I I I I I T n 1 1 0.00 y.OO 8.00 12.00 16. 20.00 24.00 28.00 TIME (MINUTES) (xIO* I Figure D.IO. Plot of observed values and the predicted lines for PAH concentration vs time for AC-1. After a 750 mg primer dose of aPAH, 36.9 mg aPAH/min was infused. The equation of the predicted line is PAH concentration = (1.56 x 10 x time + 0.0029. OBSERVED PAH t\j PREDICTED LINE o -Jo CO LU O z' 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) (xio* i Figure D.ll. Plot of observed values and the predicted line for aPAH concentration vs time for experiment AC-2, performed on steer #8232 on November 18, 1980. The animal weighed 320 lbs. and received a 725 mg primer dose of aPAH and a continuous infusion of 36.4 mg aPAH/min. The equation of the predicted line is aPAH concentration = (4.27 x 10 x time + 0.016. AC 2 OBSERVED RPfiH PREDICTED LINE O I—I n 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) (xio' i Figure D.12. Plot of observed values and the predicted lines for PAH concentration vs time for experiment AC-2. After a 725 mg primer dose of aPAH, 36.4 mg aPAH/min was infused. The equation of the predicted line is PAH concentration = (6.15 x 10 x time + 0.056. AC 2 OBSERVED PfiH PREDICTED LINE D *—I 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) txio^ i Figure D.13. Plot of observed values and the predicted line for aPAH concentration vs time for experiment AC-3, performed on steer #8241 on November 18, 1980. The animal weighed 338 lbs. and received 770 mg primer dose of aPAH and a continuous infusion of 38.4 mg aPAH/min. The equation of the predicted line is aPAH concentration = (1.62 x 10-4 ) x time + 0.003. AC 3 a OBSERVED fiPRH PREDICTED LINE n 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) cxio' i Figure D.14. Plot of observed values and the predicted lines for PAH concentration vs time for experiment AC-3. After a 770 mg primer dose of aPAH, 38.4 mg aPAH/min was infused. The equation of the predicted line is -4 PAH concentration = (2.61 x 10 ) x time + 0.04. AC 3 OBSERVED PAH PREDICTED LINE CO UJ n 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) (xio' i Figure D.15. Plot of observed values and the predicted line for aPAH concentration vs time for experiment AC-4, performed on steer #8241 on November 20, 1980. The animal weighed 338 lbs. and received 770 mg primer dose of aPAH and a continuous infusion of 38.4 mg aPAH/min. The equation of the predicted line is _5 aPAH concentration = (-2.83 x 10 ) x time + 0.44. a AC 4 o OBSERVED RPRH PREDICTED LINE Zo \10 a* n LJO o o 1 1 1 1 1 1 1 0.00 y.OO 8.03 12.00 16.00 20.00 24.00 28.00 TIME (MINUTES) txio' j Figure D.16. Plot of observed values and the predicted lines for PAH concentration vs time for experiment AC-4. After a 770 mg primer dose of aPAH, 38.4 mg aPAH/min was infused. The equation of the predicted line is PAH concentration = (-2.89 x 10 x time + 0.026. AC Ik OBSERVED PAH PREDICTED LINE CO UJ I—I n 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) (xio' i Figure D.17. Plot of observed values and the predicted line for aPAH concentration vs time for experiment AC-5, performed on steer #8263 on December 5, 1980. The animal weighed 355 lbs. and received 805 mg primer dose of aPAH and a continuous infusion of 40.3 rag aPAH/min. The equation of the predicted line is aPAH concentration = (5.06 x 10 x time - (2.98 x 10 ^). AC 5 o OBSERVED fiPRH PREDICTED LINE a n 0- 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) cxio^ j Figure D.18. Plot of observed values and the predicted lines for PAH concentration vs time for experiment AC-5. After a 805 mg primer dose of aPAH, 40.3 mg aPAH/min was infused. The equation of the predicted line is PAH concentration = (-1.70 x 10 x time + 0.039. o AC 5 OBSERVED PAH PREDICTED LINE CO UJ o 0.00 8.00 12.00 16.00 20.00 28.00 TIME (MINUTES) (xio» j