DEVELOPMENTAL ASPECTS OF PIGMENTATION IN THE MEXICAN LEAF FROG, PACHYMEDUSA DACNICOLOR
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Authors Frost, Sally Kay Viparina
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University Microfilms International 300 North Zeeb Road Ann Arbor, Michigan 48106 USA St John's Road, Tyler's Green High Wycombe, Bucks, England HP10 8HR FROST, 8ALLY KAY VXPARZNA "DEVELOPMENTAL AS^CTS OF PX6MENTATX0N XN THE HEXXCAN LEAF FROft; PACHYMlDUIA 0ACNXCOLOR, ~
THE UNZVER8XTY OF ARZZONA* PH?Dt, 1976
University Microfilms International 300 N Z£EB ROAD, ANN ARBOR. Ml 48106 DEVELOPMENTAL ASPECTS OF PIGMENTATION
IN THE MEXICAN LEAF FROG,
PACHYMEDUSA DACNICOLOR
by
Sally Kay Viparina Frost
A Dissertation Submitted to the Faculty of the
DEPARTMENT OF CELLULAR AND DEVELOPMENTAL BIOLOGY
In Partial Fulfillment of the Requirements For the Degree of
DOCTOR OF PHILOSOPHY WITH A MAJOR IN BIOLOGY
In the Graduate College
THE UNIVERSITY OF ARIZONA
19 7 8 THE UNIVERSITY OF ARIZONA
GRADUATE COLLEGE
I hereby recomnend that this dissertation prepared under my direction by Sally Kay Viparina Frost entitled Developmental Aspects of Pigmentation in the Mexican
Leaf Frog, Pachymedusa dacnicolor be accepted as fulfilling the dissertation requirement for the degree of Doctor of Philosophy
erlatlon Director Date
As members of the Final Examination Committee, we certify that we have read this dissertation and agree that it may be presented for final defense. 72? /??& A?Insu: L /U3 , y l •Wcl U 7r <2$ eypu^^/A., 11?-? ,-ZLJb / /*??
Final approval and acceptance of this dissertation is contingent on the candidate's adequate performance and defense thereof at the final oral examination. STATEMENT BY AUTHOR
This dissertation has been submitted in partial fulfillment of requirements for an advanced degree at The University of Arizona and is deposited in the University Library to be made available to bor rowers under rules of the Library.
Brief quotations from this dissertation are allowable without special permission, provided that accurate acknowledgment of source is made. Requests for permission for extended quotation from or re production of this manuscript in whole or in part may be granted by the head of the major department or the Dean of the Graduate College when in his judgment the proposed use of the material is in the in terests of scholarship. In all other instances, however, permission must be obtained from the author.
SIGNED ACKNOWLEDGEMENTS
I would like to take this opportunity to thank a number of people for their support, cooperation and participation during these studies. All of the members of my committee have been extremely help ful. Dr. Joseph T. Bagnara, aside from his helpful suggestions and criticisms as my dissertation director and major professor, has been both supportive and encouraging throughout these studies. His blind faith in my abilities and his personal warmth and sympathy throughout my graduate years were largely responsible for my continuing interest in pigment cell biology and amphibian development and for the accom plishment of this dissertation. Similarly, I would like to express my thanks to Dr. Mac Hadley for both his encouragement and sympathetic advice during the past four years. His confidence in my ability as a researcher has always been a stabilizing factor to my often shaky ego.
To both of these men I owe a sincere debt of gratitude.
The other members of my committee, Dr. Wayne Ferris, Dr. Konrad
Keck and Dr. Walter Miller have also been continually helpful and sup portive. I am grateful to Dr. Ferris for his patience and cooperation in doing the electron microscopy for some peripherally related studies.
I appreciate the time and advice Dr. Keck has offered and regret that I have not spent more time tapping his "vast storehouse of knowledge".
Lastly, I can not thank Dr. Miller enough both for putting up with me
iii as a T.A. and then consenting to substitute on my committee only to willingly fill in permanently.
Bryan Fuller, fellow graduate student and cohort, has always been helpful both with suggestions and in finding or lending equipment
(especially someone else's). A number of students involved either directly or indirectly also should be thanked. These include Anita
Switzer, Judy Frank, Cindy Stackhouse and Frank Tomizuka.
Finally, and foremost, I would like to thank my husband John.
Apart from his emotional support including confidence-building, he was helpful throughout these studies. His critical comments of my rough drafts of this dissertation, his help in collecting animals in Mexico, his photographic skills and his patience and understanding during the most difficult times of the past two years have contributed greatly to my well-being and to the completion of this degree. TABLE OF CONTENTS
Page
LIST OF ILLUSTRATIONS vii
LIST OF TABLES ix
ABSTRACT x
1. INTRODUCTION 1
Pteridines 2 Biochemistry of Pteridines 3 Metabolism of Pteridines 6 Occurrence of Pteridines 13 Xanthine Dehydrogenase 14 Avian XDH 16 Mammalian XO 19 Drosophila XDH 22 Pachymedusa dacnicolor 25 Description and Natural History of Pachymedusa . . 25 Physiology of Pachymedusa 26 Pigment Biology of Pachymedusa 27 Developmental Biology of Pachymedusa 30 Statement of the Problem 31
2. MATERIALS AND METHODS 33
Maintenance and Rearing of Animals 33 Adults 33 Embryos and Larvae 34 Chemicals 35 Experimental Procedures 36 Thyroxine Implants 36 Pteridlne Extraction and Identification .... 37 Xanthine Dehydrogenase Studies 44
3. RESULTS 49
Thyroxine Experiments 49 Pteridine Analysis 50 Adult Pteridines 52 Larval Pteridines . 57
v vl
TABLE OF CONTENTS—Continued
Page
Xanthine Dehydrogenase Studies 65 Extraction, Characterization and Assay of XDH . . 65 Inhibition of XDH 73
4. DISCUSSION 80
Thyroxine and Pterorhodin Deposition 80 Pterorhodin, Pteridine Precursors and Leaf Frog Development 83 Xanthine Dehydrogenase and Leaf Frog Pigmentation . . 93 Summary 97
REFERENCES CITED 101 LIST OF ILLUSTRATIONS
Figure Page
1. The Structural Basis of Pteridines 4
2. Probable Metabolic Pathways for the Unconjugated Pteridines 10
3. Reactions Catalyzed by Xanthine Dehydrogenase 17
4. Local Effects of Thyroxine on Pachymedusa dacnicolor Skin 51
5. Dowex 1X8 (Formate) Column Chromatography of an Adult Pachymedusa dacnicolor Skin Extract 53
6. Column Chromatographic Purification of Two Pteridines from Pachymedusa dacnicolor Skin 54
7. Absorption Spectra of Two Purified Pteridines 56
8. Two-Dimensional Thin Layer Chromatography of an Early Larval Pachymedusa dacnicolor Skin Extract . . .59
9. Thin Layer Chromatogram of Skin Pteridines from Representative Stages of Pachymedusa dacnicolor . . .61
10. Electrophoretic Analysis of Xanthine Dehydrogenase from Various Tissues of Pachymedusa dacnicolor ... 66
11. Pteridine Conversion by Pachymedusa dacnicolor Xanthine Dehydrogenase 68
12. Developmental Fluctuations in Leaf Frog Liver XDH .... 70
13. Developmental Fluctuations in Leaf Frog Kidney and Skin XDH . 71
14. Thin Layer Chromatographic Comparison of Early Larval Control vs. Allopurinol-Treated Pachymedusa dacnicolor Skin Extracts 76
vii viii
LIST OF ILLUSTRATIONS—Continued
Figure Page
15. Alterations between Control and Allopurinol-Treated Skins during Pachymedusa dacnicolor Development ... 78
16. Pteridines Believed to be Involved in the Synthesis of Pterorhodin 87 LIST OF TABLES
Table Page
I. Enzymes Involved in the Metabolism of Unconjugated Pteridines . 8
II. Physico-Chemical Properties of Selected Pteridines .... 15
III. Physico-Chemical Properties of the Pteridines Isolated and Purified from Pachymedusa dacnicolor Skin .... 58
IV. Developmental Record of Alkaline Skin Extracts from Leaf Frogs 62
V. Developmental Record of Ethanol Skin Extracts from Leaf Frogs 63
VI. XDH Activity from the Skin and Serum of Metamorphosing Pachymedusa dacnicolor as Determined from Crude Tissue Extracts 72
VII. The Inhibition of Xanthine Dehydrogenase Activity during Pachymedusa dacnicolor Development as Determined by Partially Purified Enzyme Extracts ... 74
VIII. The Inhibition of Xanthine Dehydrogenase Activity during Pachymedusa dacnicolor Development as Determined by Crude Enzyme Extracts 75
ix ABSTRACT
A study of some of the developmental aspects of pigmentation in
Mexican leaf frogs, Pachymedusa dacnicolor, was undertaken. Emphasis was placed primarily on the pteridine pigments, and in particular, pterorhodin. Pterorhodin, a red pteridine dimer, is the predominant pigment of adult leaf frog skin but is not present in the skins of lar val leaf frogs. This red pigment makes its first appearance at the very end of metamorphosis. Unlike other pteridines which are generally associated with pterinosomes, pterorhodin is a melanosomal pigment, occurring in association with melanin.
Questions concerning the biochemical and hormonal events leading up to the synthesis and deposition of pterorhodin were asked. The iso lation and identification of the pteridines occurring throughout development in the skins of leaf frogs was accomplished. From results in this study it seems likely that 7-methyl-xanthopterin, which appears in skins at the beginning of metamorphosis, and xanthopterin, which is present throughout development but increases quantitatively at the beginning of metamorphosis, are the natural precursors to pterorhodin.
Leucopterin which appears at the same developmental time as pterorhodin is probably an artifact of oxidative degradation of pterorhodin. Until this report, 7-methyl-xanthopterin, leucopterin and pterorhodin had not been reported to occur in a vertebrate.
x xi
A role for the enzyme, xanthine dehydrogenase, in leaf frog pteridine synthesis was established. Use of the xanthine dehydrogenase inhibitor, allopurinol, on leaf frog tadpoles demonstrated decreases in certain skin pteridines including a retardation in the appearance of pterorhodin at the end of metamorphosis. Superficial properties of the enzyme, including its appearance in sera of developing larvae and its dual functional role in excretion (uric acid production) and pigmenta tion (pteridine pigment production) are discussed.
Speculation concerning the synthesis, localization and turnover of pteridines is presented. The developmental role of thyroxine in the deposition of pterorhodin and in the possible regulation of xanthine dehydrogenase is considered. Finally, the occurrence of xanthine dehy drogenase and pteridines in the serum is noted in a discussion of the possibility of enzyme and/or pigment transport and deposition. CHAPTER 1
INTRODUCTION
An introduction to the study of pigmentation of the Mexican
leaf frog, Pachymedusa dacnicolor, must include a discussion of several
topics which might at first glance seem to be rather unrelated. In
cluded in these and of primary significance to the understanding of
pigment development in this organism is a basic knowledge of a group of
pigments known as pteridines. While not the only pigments present in
the skin of these animals, pteridines predominate in both quantity and
variety. Moreover, they undergo a number of developmental changes of
importance to this particular organism and with broad significance to
more fundamental problems such as eukaryotic regulatory mechanisms and
the control of gene expression during development. Secondarily, empha
sis was placed on a particular enzyme, xanthine dehydrogenase. This
enzyme has several functions and has been studied in a number of systems. Unfortunately, little is known about the amphibian enzyme,
and data from a variety of other sources were used to extrapolate
information pertinent to Pachymedusa dacnicolor. An understanding of
the functions of xanthine dehydrogenase and the properties associated with this enzyme are necessary in order to comprehend the evolutionary
and biochemical roles of its end products, uric acid and pteridine
pigments. Finally, amphibians as a developmental tool are briefly
1 2 discussed. What little is known about the physiology, biochemistry and pigment development of Pachymedusa is considered in relation to the present study. Taken together, this information provides the founda tion for the work discussed in the subsequent pages.
Pteridines
The importance of pteridines as biological compounds is well known. Examples of their ubiquitous occurrence abound in the litera ture and such occurrences vary from their participation as co-factors in a number of biological reactions (Kaufman, 1967), to their inhibi tion of RNA and DNA synthesis in insects (Harris and Forrest, 1967;
Puckett and Snyder, 1975; Smith and Forrest, 1976).
Of significance to the present study is the function that pter idines are known to have in the pigmentation of lower vertebrates
(Bagnara, 1961; Hama, 1963; Hama and Fukuda, 1964; Obika and Bagnara,
1964), and invertebrates (Ziegler and Harmsen, 1969). Of note are the white, yellow and orange colors found in butterfly wings (Pfleiderer,
1964), the red and yellow insect eye pigments (Kaufman, 1967), and the red and orange coloration of the dewlap of chameleons (Ortiz and
Williams-Ashman, 1963). Unfortunately, with the exception of some yellow colored pteridines, such as sepiapterin and xanthopterin, and the few orange and red pteridines, like the drosopterins and ptero- rhodin, most of the naturally occurring pteridines are colorless
(Gates, 1947). Thus, their role as pigments is somewhat questionable, even though they are clearly deposited in discreet pigment cell organ elles such as those found in vertebrate skin (Matsumoto, 1965). 3
With respect to pigment cell biology, it is perhaps unfortunate that much of the emphasis on the biochemistry of pteridines was placed on the biological role of folic acid derivatives (Blakley, 1969).
Micro-organisms provided the tool that allowed the initial genetic and biochemical studies on pteridine metabolism to be carried out (Brown,
1971). Even now, experiments employing prokaryotic systems serve as a basis for similar studies in insects and vertebrates. Fortunately, the compounds themselves, as well as what is known about the metabolism of these compounds, appears to have been phylogenetically conserved
(Kaufman, 1967). In general, the enzymes from organism to organism are all functionally alike but may differ in fundamental structure such that they are electrophoretically or immunochemically distinct. Thus, while the initial step in the synthesis of all pteridines is the same, the pathway diverges early along. Beyond the initial steps much less is known about the enzymes and mechanisms involved in the conversions of those pteridines functioning as pigments than is known about folic acid-related compounds.
Biochemistry of Pteridines
Chemically, pteridines can be grouped into two broad cate gories: (1) the "unconjugated" or simple pteridines, and (2) the
"conjugated" pteridines. The first group includes xanthopterin, leucopterin and all of the other naturally occurring pteridines pre sumably involved in pigmentation. With one exception all possess the basic "pterin" structure shown in Figure 1, and contain an -NH2 and =0 group in the 2 and 4 positions, respectively. The exceptions to this 4
4 5 r>
i 8 pteridine ring system
"C^jO (pterin) unconjugated pteridine
COOH
Hfjl' jp ^|CH2-NH C-NH-CH CH« H-N " ch2 (folic acid) COOH conjugated pteridine
Figure 1. The Structural Basis of Pteridines. 5
are the lumazines which contain =0 groups at both positions. Conju
gated pteridines are those of the folic acid family, and all contain
either £-aminobenzoic acid or £-aminobenzoylglutamic acid as a sub-
stituent. All pteridines are now known to be synthesized from a common
natural precursor, guanosine triphosphate (Rembold and Gyure, 1972).
Excellent reviews on the chemistry of folic acid and related pteridines
are presented by Blakley (1969), Ziegler and Harmsen (1969) and Brown
(1971).
No discussion of the simple pteridines is complete without
mention of some of their distinctive chemical properties. Pteridines,
like their close relatives, the purines, are generally insoluble at
neutral pH. Most pteridines can be solubilized in dilute alkali or
acid, but care must be exercised that such conditions, alkali in par
ticular, will not cause either the breakdown or conversion of the
particular pteridine. In addition to alkaline degradation, many pteri dines are extremely photolabile and must be handled under reduced light or in total darkness. A red "safe light", commonly found in photo
graphic darkrooms, is suitable for such work. Care must also be taken to prevent oxidative breakdown of some pteridines, and 2-mercapto- ethanol (.01%) is generally included in extracts and solvents for this
purpose.
Apart from the above mentioned problems encountered in working with pteridines, there are several advantages. Most simple pteridines fluoresce when exposed to ultraviolet light, and in many cases the fluorescent color alone is an indicator of a particular pteridine or group of pteridines. These small molecular weight compounds are easily 6
separated by paper or thin layer chromatography (TLC) and can be
quantitated using a fluorometer. Generally, pteridines are identified
by at least three criteria: (1) Rf values from TLC, paper chroma
tography and/or paper electrophoresis in several different solvents;
(2) fluorescent color; and (3) absorption and/or fluorescent emission
spectra recorded at appropriate wavelengths. These three criteria when
compared with the same information obtained from authentic, pure
pteridines, are generally sufficient to accurately identify any pteri-
dine. In a few cases pteridines even possess a very distinctive
visible color, such as xanthopterin which is yellow in solution, or
pterorhodin which is red in acid solutions, purple at neutral pH, and
rapidly converts from red to orange to yellow in alkali. The great
majority of simple pteridines are, however, clear in solution, and a
few, such as the tetrahydropteridines, do not even fluoresce (Gates,
1947; Pfleiderer, 1964).
Metabolism of Pteridines
For many years the question of pteridine precursors was a
mystery with most workers agreeing that the likely candidate was a
purine or a purine derivative (Albert, 1957). With the advent of radiolabelling techniques it was a relatively simple matter to demon
strate that pteridines were synthesized most efficiently when Re labelled guanosine triphosphate was the precursor supplied (Shiota and
Palumbo, 1965). Moreover, it has now been shown in a variety of sys
tems that guanine is the purine of choice, and the ribose component of 7 the nucleotide as well as the triphosphate are essential for the ex pression of maximal pteridine synthesis (Brown, 1971).
The specificity for guanosine triphosphate is determined by the requirements of GTP cyclohydrolase, the first enzyme in the chain responsible for the production of pteridines from GTP. While GTP cyclohydrolase has been isolated and characterized from a number of systems (see Table I), Rembold and Gyure (1972) insist that this is probably not the only pathway initiating pteridine biosynthesis since at least two forms of GTP cyclohydrolase have been reported to exist.
However, Foor and Brown (1975) have recently completed a detailed com parison of the properties of GTP cyclohydrolase I and II from Escheri chia coli. They have concluded that GTP cyclohydrolase II is most likely involved in the biosynthesis of riboflavin, while I most certainly synthesizes the precursors to folic acid and unconjugated pteridines. So, at least for now, the evidence suggests that all pteridines and pteridine derivatives originate from the immediate product of GTP cyclohydrolase action.
Figure 2 summarizes what is known about the metabolism of sim ple pteridines. This pathway was constructed primarily from what is known to occur in micro-organisms and what has been found to occur in insects and, to a lesser extent, in vertebrates. An extensive list of references is provided with Figure 2, and some emphasis should be placed on understanding pteridine metabolic interrelationships since a large part of the discussion to follow concerns these pathways.
Studies on the isolation, identification and chemical synthesis of pteridines were primarily accomplished during the late 1940's and Table I. Enzymes Involved in the Metabolism of Unconjugated Pteridines.
Enzyme Source M.W. pll Opt. Km Cofactor Miscellaneous Properties References
GTP Streptomyces 500,000 8 20 nH none requires EDTA for full activity; O2, Elstner & Cyclohydrolase rlmosus (GTP) PCMB and shodonycln are Inhibitors; Suhadolnik, 1971 ATP Is a competitive Inhibitor +2 Comamonas 650,000 7.6 3 uM none Mg Is necessary for activity Cone & Guroff, 1971 sp. (GTP) Cone, Plowman & Curoff, 1974
Escherichia 210,000 8 22 nH none divalent metal Ions (except Hg+2 ) are Burg & Brown, 1968 coll (GTP) Inhibitors; ATP dCTP and guanoslne-5'- Brown, 1971 (I) tetraphosphate are competitive Inhibi Ylm & Brown, 1976 tors; synthesizes folic acid, etc. +2 Escherichia 44,000 8.4 41 uM none activity Is stimulated by Mg ; Brown et al., 1975 coll (CTP) Inhibited by EDTA and Inorganic phos Foor & Brown, 1975 (11) phate; synthesizes riboflavin
Drosophlla 345,000 7.8 22 uM none divalent cations are Inhibitory Brown & Fan, 1975 melanoeaster (GTP) Fan & Brown, 1976
Chicken 125,000 8-9.2 14 uM none Cu*2 and Hg"^ are inhibitors; lmnuno- Fukushima, Richter Liver (GTP) loelcally distinct from E. coll enzyme & Shlota, 1977 lU-Neopterin Escherichia 87-89,000 6 ? none activity is stimulated by Hg"*"2 or Mn^ Brown et al., 1975 Eplmerase coll llj-Neopterln- Escherichia 17,000 8.5 11 uM none Mg+^ is necessary for activity; Suzuki & Brown, (P)j Pyrophos- coll (HP) substrate decomposes in the presence 1974 pliohydrolase of Mn lU-Neopterin Escherichia 100,000 9.6 9 uM none end product is a potent inhibitor; Mathls & Brown, Aldolase coll (HP) EDTA Increases enzyme activity 1970
ClljOII-l^-Pteri- Escherichia 15,000 8.5 .15 uM none Mg+2 Is necessary for activity Rlchey & Brown, dine Pyrophos- coll 1971 phoklnase Table I. - (Cont'd.)
Enzyme Source M.W. pH Opt Km Cofactor Miscellaneous Properties References
Seplapterin Bombyx 8 59 mM none A1IP, XP and BP are all Inhibitory; Tsusue, 1971 Deaminase mori (SP) biological role is unknown
Seplapterln rat liver 5.5 .41 uM NADH or £-mercurlbenzoate or 2,4-DNP are Matsubara et al., Reductase (SP) NADPI1 Inhibitors 1966
horse liver. 47 000 5.5 .21 uM NADll or j>-mercuribcnzoate, 2,4-DNP and various Katoh, 1971 (SP) NADPII unconjugated ptcrldlnes are inhibitors
Pterin AlcallRenes 6.5 ? none PCMB and KF are inhibitors; specific Levenberg & Deaminase metacallgenes for un- or 6-substltuted pterldines ilayashl, 1959
rat liver 6.5 30 mM none IXP and H -pterin are also deamlnated; Rembold & (A1IP) azaguanlne and lumazlne are inhibitors Simmersbach, 1969
Xanthine Clostridium 1 .16 uM HAD requires FAD, molybdenum and iron for Bradshaw & Barker, Dehydrogenase cyllndrosporum (xanthine) activity 1960
Bombyx 8.4 ? NAD requires FAD, molybdenum and Iron for ilayashl, 1961 morl activity
Drosophila 300 000 ? .24 uM NAD requires FAD, molybdenum and iron for Parzen & Fox, 1964 melanoKaster (xanthine) activity; Andres, 1976
Chicken 300 000 7.5 .33 uM NAD requires FAD, molybdenum and iron for Strittmatter, 1965 Liver (xanthine) activity; adenine, azaxanthlne and Rajagopalan & guanine are Inhibitors Handler, 1967
Bovine 300 000 7.8 ? NAD requires FAD, molebdenum and Iron for Nelson & Handler, Milk activity 1968
Isoxanthopterin Bombyx 6.6 .12 uM none extremely specific for IXP; XP, Gyure, 1974 Deaminase morl (IXP) quanlne, KF and KCN are inhibitors
Erytliropterin Oncopeltus 7.5 1 mM 7 requires xanthopterin and oxalo- Forrest, Menaker & Synthetase fasclatus (XP) acetate for activity Alexander, 1966 Figure 2. Probable Metabolic Pathways for the Unconjugated Pteridines.
This illustration was constructed from information derived from the following sources: Brown, 1971; Brown and Fan, 1975; Brown et al., 1975; Eto, Fukushima and Shiota, 1976; Forrest, Menaker and Alexander, 1966; Fukushima, 1970; Fukushima and Shiota, 1974; Goto and Suguira, 1971; Gyure, 1974; Harmsen, 1970; Krumdieck, Shaw and Baugh, 1966; Levy, 1964; Needham, 1974; Okada and Goto, 1965; Parisi, Carfagna and D'Amora, 1976; Pfleiderer, 1964; Rembold and Gyure, 1972; Richey and Brown, 1971; Shiota, 1971; Stackhouse, 1966; Suguira and Goto, 1968; Suzuki and Brown, 1974; Watt, 1967; Ziegler and Harmsen, 1969. PLEASE NOTE:
Print on some pages is small and indistinct. Filmed in the best possible way.
UNIVERSITY MICROFILMS. otp cyclohtoaolasc sx c^0m3 mj-ncopterin Co j£0 «i L-fhr«oH.-NEOPTERIN-(P), GTP O-crythro H2- NEOPTERW-fMj PP. ^4 «2 NEOPfERUMPIj f PYROftttSPHOHVOftOlASCpyi \ O-trylhro-Hj -NEOPTERIN- P -vs&s.^ t 9*9" ch*ch'chaoh AAA — 'l h^-biopterin biopterin i 'i SEPlAPTtftIN REDUCTASE O-^yihfo-Hg-NEOPTERINz / ' hj-ncoptehln f f ch scpiaptchin hn' is-ch, raldolase | / i IK- a chjoh- h2- o • / t *2* °k N chJipi2 pyridine h folic acid SEPIAPTERIN hj-«- lactdtl- lumazine PTROPHOSPHO- MjjWkx.j.. / S Otfi 'OhtfCS SWTMETASE KINASE j clljohhj-pterichne ' » drosoptemns w K "VT '' y'^v'v g * ^|0h c'">h y^y"^ 1 mf*if'"5*]
sr n jCCc XXX ijO £QH CO leucopterin CHRYSOPTERIN ERYTHROPTERIN LUMAZINE 7-oxylumazine
v ....4... J
J hg « ^«Hj pterorhooin a other C-7 ttibalilultd pltridlMt
Figure 2. Probable Metabolic Pathways for the Unconjugated Pteridines. o throughout the 1950's. Once the chemistry of these compounds was
established, emphasis turned towards elucidating the natural mechanisms
for pteridine synthesis. Again, radiolabelling techniques were instru
mental in the experiments designed to clarify pteridine metabolic
interrelationships. However, even today, after nearly 30 years of
sporadic work on pteridine biosynthesis in a variety of systems, the
state of what is known as shown in Figure 2 is still somewhat chaotic.
For example, until recently sepiapterin was believed to be the
precursor to biopterin. This idea was supported by the discovery of
the enzyme sepiapterin reductase (Matsubara et al., 1966; Katoh,
1971), which reversibly catalyzes the conversion of sepiapterin to di-
hydrobiopterin. The reversibility of this reaction was initially
overlooked and the logical conclusion was that sepiapterin preceded
biopterin in the metabolic pathway. Only recently were Parisi et al.
(1976) able to demonstrate convincingly that, at least in Drosophila,
biopterin must precede sepiapterin.
A closer look at Figure 2 reveals four branching pathways, two of which are significant to this study, one of which is of minor im portance and one which will not be considered here. The pathway leading to folic acid is not important to this discussion and is included in Figure 2 simply to demonstrate the relationship of folic acid to unconjugated pteridines. The biopterin pathway is significant only as far as biopterin itself is concerned. Most of the emphasis in this study has been placed on pathways leading to the synthesis of the
C-6 and C-7 substituted pteridines, and in particular to pterorhodin. Unfortunately, very little is known about the synthesis of either the
C-6 or C-7 substituted pteridines, and even less is known about the
synthesis of pteridine dimers such as pterorhodin and the drosopterins.
Many of the oxidative reactions leading to the synthesis of pterorhodin
are believed to occur nonenzymatically (Needham, 1974). However, these
reactions generally occur at unreasonably low pH's, increasing the
likelihood for the existence of enzymatic mechanisms as yet unidenti
fied.
It is fortuitous that pteridines function as eye pigments in
Drosophila, as well as being found throughout the bodies of these
insects (Ziegler and Harmsen, 1969). As eye pigments, they have been
subjected to the elegant genetic studies commonly associated with
Drosophila (Ziegler, 1961). While such studies contributed considera
bly to what is known of the sequence of events in pteridine synthesis,
they were less helpful in determining which events were enzymatically
controlled and what kinds of enzymes were involved.
The most terminal of the enzymes known to interconvert pteri
dines is xanthine dehydrogenase (XDH). Beyond this, pteridine inter-
conversions continue with little information as to how these intercon-
versions are accomplished or how pteridines are finally catabolized.
Pteridine catabolism is believed to proceed in two directions. First,
the discovery of a number of deaminases demonstrates that many pteri
dines are catabolized to lumazines (Fig. 2). Secondly, pterin-6-
carboxylic acid is known to arise as a result of folic acid catabolism
(Krumdieck et al., 1966). It is also suspected to arise from the 13 oxidative degradation of biopterin and C-6 substituted pteridines.
As a result, pterin-6-carboxylic acid is not shown in Figure 2, primarily because it arises from such a variety of sources, and its natural synthesis, if it has one, has not been clearly established.
In summary, what little is known about the enzymes involved in the synthesis of simple pteridines is outlined back in Table I. From this table it is readily apparent that the bulk of enzymatic informa tion is derived from 12. coli and other micro-organisms. Moreover, most of this work has been accomplished just since 1970, and the trend now is to verify these enzymatic mechanisms in other organisms and proceed to identify other enzymes. To date only four of these enzymes have been identified in vertebrates, and one of these (XDH) was identified for reasons not originally associated with pteridine synthesis.
Occurrence of Pteridines
Much of what we now know concerning pteridine interconversions came from studying organisms that are rich in at least several of these compounds. Thus, insects, fish, frogs and lizards have been the sys tems of choice, and some interesting generalizations have been made concerning the nature and kinds of pteridines found in these organisms.
In general, insects are known to exhibit pteridines which occur along all three of the pathways leading to the synthesis of unconju gated pteridines as illustrated in Figure 2 (Ziegler and Harmsen,
1969). As a result some pigments, such as leucopterin, chrysopterin
(7-methyl-xanthopterin), and pterorhodin, were associated only with certain insects. In vertebrates, the pathway leading to isoxantho- pterin is believed to predominate, and xanthopterin, while often present in small amounts, is not considered a dominant vertebrate pteridine. Those pteridines following xanthopterin in the metabolic chain of C-7 substituted pteridines are not reported to occur at all in vertebrates.
Drosopterins, although named for their original source,
Drosophila, have been commonly found in insects and vertebrates alike, and are generally associated with red and red-orange coloration.
Chemically elusive compounds, drosopterins have only recently been identified as pteridine dimers (Pfleiderer, 1970), and as such, they are chemically related to another red pteridine dimer, pterorhodin
(Russell et al., 1949). These dimers all share several distinct properties: (1) extreme insolubility in anything except harshly acidic or alkaline solutions, (2) low Rf values in organic solvents, and (3) absorption maxima extending above 400 nm.
In summary, those pteridines significant to the present study are listed in Table II. The accompanying information is supplied as a convenient reference for the work outlined herein.
Xanthine Dehydrogenase
Of particular interest to the present study is the enzyme, xanthine dehydrogenase (xanthine:oxygen oxidoreductase, E.C. 1.2.3.2.).
Much is known about this enzyme in a number of systems, but little information is available concerning either the occurrence, properties or role of xanthine dehydrogenase in amphibians. Studies on xanthine Table II. Physico-Chemical Properties of Selected Pteridlnes. Natural Fluores. Compound Color Absorption Spectra, Xmax Color General Properties References
Leucoptcrln Colorless 240,282,342 (In 0.1 N NaOH) Bright Insoluble In dilute acids and Jacobson & Simpson, (LP) Blue water; soluble In dilute alkali, 1946 conc. H2SO4; occurrence limited Gates, 1947 to insects
Xanthopterin Yellow- 256,392 (in 0.1 N NaOH) Bright slightly soluble In water; Jacobson & Simpson, (XP) Green 230,261,356 (In 0.1 N HC1) Blue- soluble In dilute acids and 1946 Green alkali; widespread occurrence Ellon & Hltchlngs, from insects to humans 1947
Isoxanthopterin Colorless 225,255,340 (In 0.1 N NaOH) Violet considerably more acidic than Ellon & Hltchlngs, (IXP) 287,340 (in 0.1 N HC1) XP; relatively insoluble in 1947 acids; widespread occurrence Goto & Suguira, 1971
D-erythro-Neopterin Colorless 220,255,365 (in 0.1 N NaOH) Blue somewhat soluble in water; Coto & Suguira, 1971 (HP) 248,322 (in 0.1 N HC1) readily soluble in acid and Rembold & Gyure, alkali; widespread occurrence 1972
7-Methyl- Yellow- 253,344 (in 0.1 N NaOH) Blue- Insoluble in organic solvents Ellon & Hltchlngs, Xantliopterin Green 232,265,359 (in 0.1 N HC1) Green & water; soluble in dilute acid 1947 (7-CH3-XP) and alkali; occurrence limited Pfleiderer, 1964 to Insects
Pterin Colorless 253,362 (in 0.1 N NaOH) Blue soluble in dilute acid and Pfleiderer, 1964 (AIIP) 243,313 (in 0.1 N 1IC1) alkali; somewhat soluble in Coto & Suguira, 1971 water; widespread occurrence
Pturin-6- Colorless 263,364 (in 0.1 N NaOH) Blue soluble in dilute alkali; Pfleiderer, 1964 Carhoxylic Acid 260,348 (in 0.1 N HC1) somewhat soluble in dilute Goto & Suguira, 1971 (AHP-6-C001I) acid & water; widespread occurrence
Uiopterln Colorless 254,363 (in 0.1 N NaOH) Blue soluble in acid, alkali and Pfleiderer, 1964 (BP) 263,322 (in 0.1 N HC1) organic solvents; somewhat Goto & Suguira, 1971 soluble in water; widespread occurrence
Pterorhodin Red 325,360,508,535 (in conc. Deep insoluble in dilute acid, Russell et al., (Pt) H2SO4) Blue- organic solvents and water; 1949 240,325,502,540 (in 88Z Violet soluble in strong acids; Pfleiderer, 1970 formic acid) soluble in alkali, but Misuraca et al., degrades with time; occur 1977 rence limited to insects Ln dehydrogenase (XDH) or xanthine oxidase (XO) have followed several distinct approaches. While there are several differences in the physico-chemical properties between XO and XDH, the major distinction is that the oxidase utilizes molecular oxygen as its physiological electron acceptor and the dehydrogenase employs NAD in this capacity
(Rajagopalan and Handler, 1967). These enzymes, while mechanistically distinct from organism to organism, can catalyze the same reactions regardless of the system in question (Fig. 3). Most notably, XDH is responsible for the conversion of xanthine to uric acid. In addition, it also catalyzes several pteridine interconversions, such as the production of isoxanthopterin from pterin, and leucopterin from xanthopterin. A system by system discussion of this important enzyme and its properties follows.
Avian XDH
Birds in general are known to be uricotelic, excreting copious quantities of uric acid and thus requiring XDH for its synthesis
(Balinsky, 1972). Interestingly, XDH was found to have some degree of developmental significance, particularly in chick embryos. It has been shown that throughout their early development, chick embryo livers maintain a barely detectable amount of XDH. Approximately four to six hours after hatching, enzyme levels begin to rise dramatically until a maximum is reached a few days later. This maximum is maintained for several days after which it drops to a lower but constant level which is typical of chickens three months of age and older. It is remarkable that this enzyme is found in the livers of chicks just having pecked a fl h hypoxanthine xanthine uric acid
OH \^ioh
kn" "n n n pterin isoxonthopterin xanthopterin leucopterin
^cho COOH
N N' Ho-pterin l^-xanthopterin pterin-6-aldehyde pterin-6-C00H
Figure 3. Reactions Catalyzed by Xanthine Dehydrogenase. 18 hole in their shells, but is barely detectable anytime prior to the appearance of the hole (Kato, 1960).
The entire story, however, is not so straightforward. Further investigation of other embryonic chick tissues revealed XDH in the mesonephros and metanephros as well as in the liver. In the meso- nephros, XDH activity first appears during the 5th day of embryonic development. Levels continue to rise gradually until about day 12, after which enzyme activity drops off in correspondence with the resorption of the mesonephros. Simultaneously, XDH begins to appear in the developing metanephros (Croisille, 1962). It is important to note that immunochemically, electrophoretically and kinetically, the liver and kidney enzymes appear to be identical.
Finally, in studies that continued to expand the developmental aspects of chick XDH, a survey of no less than 14 different tissues added the pancreas, duodenum and intestine to the list of chick tissues with notable XDH activity. Working on the premise that since tissues like liver receive and respond to some "signal" at about the time of hatching, Fisher, Curtis and Woodward (1967) attempted to determine whether crude pituitary extracts could produce this "signal" and induce an increase in XDH activity in chick embryos. When such extracts were injected into the air sacs of embryos three days prior to hatching, a precocious accumulation of the enzyme was observed.
However, when individual pure pituitary hormones were tested alone or in varying combinations, the results could not be duplicated. Even now, it is not entirely clear which components of the crude pituitary extract were responsible for the induction of XDH activity. 19
A vast amount of work has been done on the characterization and
regulation of avian XDH. This enzyme and its inductive properties have
been used in a number of classical studies aimed at establishing whether observable increases in enzyme activity were due to actual increases in the number of enzyme molecules or, alternatively, to activation of previously inactive proteins (Murison, 1969). Regulatory studies, particularly those using steroid hormones, seem to indicate
that some sort of hormonal control mechanism exists for this enzyme
(Fisher et al., 1967). This wealth of information emphasized the
developmental significance of XDH and was a clear indication that such
an enzyme might also be of considerable importance in Pachymedusa.
Mammalian XO
The best known of the xanthine oxidases is that extracted from bovine milk. It is this enzyme that is most often available commer cially in purified form (Nelson and Handler, 1968).
In mammals, the role of xanthine oxidase is an elusive one.
In rats and mice XO is known to be present in a variety of tissues and is easily identifiable even in tissue culture. These animals also contain uricase which allows the toxic product of XO (uric acid) to be degraded. Humans, on the other hand, possess XO only in a few tissues, primarily the liver and intestinal epithelium. Human cell cultures, however, never display any detectable XO activity, and humans also have the added disadvantage of lacking uricase (Brunschede and Krooth,
1973). Cases of hyperuricemia are fairly common in humans. So, once 20 this defect was traced to a particular enzymatic disorder, the search for a suitable treatment began.
An extremely insoluble compound, uric acid, if not degraded, is deposited in the joints of the body. This painful and not uncommon affliction is generally referred to as gout (Elion et al., 1966), and the likely candidates for treatment of this metabolic disorder were at first believed to be natural inhibitors of XO. Unfortunately, natural
XO inhibitors were not effective in the treatment of gout, and a variety of substrates and synthetic analogs of substrates were tested in this capacity but with limited results (Doisy, Richert and Wester- feld, 1955). As it turned out, all substrates and a great number of analogs tested were inhibitory. Even uric acid itself was found to be a potent inhibitor of XO activity. However, with the exception of uric acid, almost all of these inhibitors were effective only when administered in unreasonably high doses and often with unwanted side effects. Using uric acid at all as a treatment would only compound the overall problem, and so another solution had to be found.
After considerably more work, an effective synthetic compound was developed. This compound is commonly known as allopurinol or HPP
(4-hydroxypyrazolo-3,4-d-pyrimidine), and was originally developed to relieve the symptoms of gout (Elion et al., 1966). Since allopurinol is of importance to the present study, it is necessary to understand some of its properties.
Allopurinol, when administered to organisms, competitively in hibits XO activity. The actual mechanism by which this inhibition 21 occurs is not a simple one. As expected, it was found that allopurinol binds the enzyme with a greater affinity (K^ for KPP being 5-40 fold
lower than that for xanthine) and faster Vmax (2-6 fold) than natural substrates such as xanthine (Spector, 1977). Moreover, this inhibition is completely reversible, although very slowly. The slowness of the reversal process turned out to be due to the tenacious binding of oxipurinol, the product of XO action on allopurinol. Thus, unlike the natural products which are rapidly released from the enzyme molecule but not readily cleared through the kidney, oxipurinol is released at a significantly slower rate but is much more readily excreted. To sus tain this inhibition, xanthine must be present (Elion et al., 1966), and reduced molybdenum must be scarce. Moreover, if an artificial electron acceptor, such as phenazine methosulfate, is substituted for oxygen, then the rapid conversion of either allopurinol or xanthine could be sustained (Spector and Johns, 1968).
Additional metabolic studies on allopurinol (Elion et al.,
1966; Alexander et al., 1966) primarily using rats and mice, demon strated that allopurinol was best administered either in the diet or by intraperitoneal injection. Moreover, the drug revealed no cumulative effects, and in order for inhibition to be maintained, a regular schedule of frequent treatment was necessary. In no case was total inhibition of XO activity ever demonstrated, nor was any inhibition of growth ever noted. Instead, allopurinol at best, significantly de creased the levels of purine biosynthesis in those organisms maintained on a steady and frequent dose of at least 30 mg/kg (body weight) of allopurinol. 22
Allopurinol has been used in a number of systems to inhibit
either xanthine oxidase or xanthine dehydrogenase (Glassman, 1965a;
Murison, 1969; Spector, 1977). As will become apparent, allopurinol is not only a treatment for gout, but a tool for the pigment cell biolo gist to approach pigmentary deficiencies as well.
Drosophila XDH
In Drosophila the emphasis on xanthine dehydrogenase is totally different from either that in birds or in mammals. Eye pigment mutants are well known in Drosophila, and one of the best documented of these occurs at the rosy (ry) locus (Glassman, 1965a; 1965b). flies have dark red-brown eyes in contrast to the normal bright red eye color of the wild type (Boni, deLerma and Parisi, 1967). Eye pigments in insects are divided into two main classes, the ommochromes and the pteridines. Both classes, while chemically distinct are believed to be metabolically reliant upon each other for proper expression of normal eye color (Parisi et al., 1976; Parisi, D'Amora and Franco, 1977).
Analysis of the pigments in r£ mutants revealed a deficiency of isoxanthopterin and an accumulation of pterin. In addition, the red drosopterins were present in markedly diminished quantities compared to that of the wild type (Glassman, 1965a; 1965b). Closer examination revealed that r%_ mutants had no detectable xanthine dehydrogenase activity. In addition, the r£ locus was definitively shown to contain the structural gene for xanthine dehydrogenase (Yen and Glassman, 1965). 23
Of particular interest were a group of studies published during the period from 1967-1971 by Parisi's group in Naples (Boni et al., 1967;
Boni and Parisi, 1967; Parisi, Figurelli and Carfagna, 1971). In these studies Drosophila larvae were exposed to various doses of allopurinol.
It was found that a concentration of .05-.08% allopurinol in media was sufficient to produce 50% phenocopies of the r^ mutant. In otherwords, in a group of wild type flies exposed to .05% allopurinol, 50% appeared to be phenotypically identical to the mutant. Biochemical investi gation of these phenocopies further confirmed the inability of XDH to function as the factor responsible for the characteristics of the ry mutant.
The extent of the work accomplished on r£ and a number of related mutants has demonstrated this complex system to be a classic one for studies in eukaryotic molecular genetics, particularly concern ing the interrelationships between control elements and structural elements. (See Chovnick, Gelbart and McCarron, 1977, for a review.)
For example, at least two other genetic loci in Drosophila are neces sary for the proper expression of the ry+ locus. One of these, lxd
Clow xanthine dehydrogenase), is believed to be the structural gene for a control element (co-factor or activator) necessary for XDH activity.
The other locus, ma-1 (maroon-like eye color), is also believed to con tain the information for a polypeptide necessary either as an activator or co-factor for XDH activity. Ma-1 mutants by themselves lack any detectable XDH and accumulate pteridines just like the r^ mutant. Lxd mutants exhibit 20-25% normal XDH activity, and the possibility of this effect being due to an inhibitor of some type has been eliminated. As a result a great deal of classical and modern genetic techniques have been employed in attempting to explain the relationships between these three loci. While jrjr is unquestionably the structural gene for XDH, the precise nature of Ixd and ma-1 is still a matter of dispute.
However, there is no question that in order for normal XDH activity to prevail, all three loci must be functioning. More details of this complex and somewhat confusing system are outlined in two excellent reviews by Glassman (1965a, 1965b).
As a final note on insect XDH, it appears that this enzyme serves the dual function of producing uric acid and pteridine pigments, both of which can, in a broad sense, be considered as excretory products (Harmsen, 1966). Moreover, while the eye itself manifests the products of XDH activity, results now confirm that the actual enzyme molecule is synthesized in a different organ and is transported to the eye (Barrett and Davidson, 1975). The evidence for this trans port phenomenon and its significance to the present study will be dis cussed in detail later.
This brief coverage of the relevant studies on xanthine dehy drogenase and xanthine oxidase can be summarized as follows: (1) At least in chick embryos, xanthine dehydrogenase is of developmental significance and is probably regulated in some fashion by hormones.
(2) This enzyire is responsible for the production of both uric acid and pteridines. While both of these functions may not be applicable to all systems, all xanthine dehydrogenases and xanthine oxidases, regardless 25
of their natural source, are capable of producing uric acid and con
verting pteridines (Krenitsky et al., 1972). (3) Finally, allopurinol
can be used to inhibit the activity of any of these enzymes, thus pro
viding a tool for the biochemical and genetic studies of XDH deficien
cies, whether pigmentary or excretory. These three points provide the
basis for studies on xanthine dehydrogenase in Pachymedusa dacnicolor.
Pachymedusa dacnicolor
No introduction to the development of pigmentation in any animal can be complete without a discussion of what is already known about the organism itself. The present study deals solely with the
Mexican leaf frog, Pachymedusa dacnicolor, a member of a family of tree frogs, the Hylidae. This particular animal is common to the western coastal plain of Mexico, from southern Sonora and south down much of the west coast of Mexico.
Description and Natural History of Pachymedusa
In appearance Pachymedusa dacnicolor is normally a striking bright green in color. However, it can assume a dark red-brown or green-gray coloration depending on a number of factors, including light, background color, and overall physiological well-being of the animal. Females of the species can grow as large as 6-10 cm, snout- vent length, and males are typically somewhat smaller in size. During most of the year, males and females are indistinguishable from each other on the basis of external appearance alone. During the breeding season, which typically extends from late June through August, males develop black callosities on their "thumbs". 26
In western Mexico, the breeding season commences with the start of the summer rains, when the daily 37°C+ temperatures are accompanied by very high humidity. Just prior to the onset of the summer rains, males are frequently found congregating at dry rain pools, presumably in anticipation of the rains. Females are seldom found in the field prior to the onset of rainy weather.
Few studies on the natural behavioral habits of this amphibian exist, apart from those by Pyburn (1970), Wiewandt (1971) and the personal field observations outlined above. Current work in this laboratory on the breeding behavior and maintenance of captive Pachy- medusa in greenhouses and other carefully regulated environments pro vided much inferential data in support of sparsely available field observations (Bagnara, Frost and Frost, in manuscript).
Physiology of Pachymedusa
In recent years hylid frogs adapted to arid environments have been of interest to physiologists and biochemists. In 1970, Loveridge noticed that Chiromantis xerampelina, a tree frog able to tolerate long periods of exposure to dry air in the African desert, excreted large amounts of a pasty white substance. Chemical analysis revealed this substance to be uric acid.
This initial discovery was of major significance since physio logically, urlcotelism was not then known and certainly not expected in amphibians. Subsequently, Shoemaker and others (Shoemaker et al.,
1972; Shoemaker and McClanahan, 1975) demonstrated uricotelism first in the South American tree frog, Phyllomedusa sauvagii, and later in other phyllomedusine frogs including Pachymedusa dacnicolor. Of all the
frogs tested and found to be uricotelic, Pachymedusa exhibited the
least tendency towards uric acid excretion, with only about 5% of its
total excretory product being composed of uric acid. Nevertheless,
the enzymatic basis for uricotelism (i.e., xanthine dehydrogenase)
was found to be prevalent in the livers and kidneys of these animals.
Moreover, little or no uricase activity was detectable in either of
these tissues, resulting in uric acid as the metabolic end product
(Vaughn Shoemaker, personal communication).
The discovery of uric acid and substantial amounts of xanthine
dehydrogenase activity in the tissues of Pachymedusa dacnicolor was of
interest to the present study, but not solely from the standpoint of
uricotelism. As will be seen, these findings are also significant in
relation to the pigmentation of this frog.
Pigment Biology of Pachymedusa
Studies on the pigmentation of Pachymedusa dacnicolor began in
1966, during initial studies on the dermal chromatophore unit of amphi
bians. Bagnara, Taylor and Hadley discovered that melanosomes in the
skin of this frog were not the same, morphologically, as all other
vertebrate melanosomes (Taylor and Bagnara, 1969). Typical vertebrate
melanosomes are uniformly electron dense, spherical to ovoid in shape,
melanin-containing, membrane-bound organelles. They are usually found
in quantity in pigment cells termed melanophores or melanocytes
(Bagnara and Hadley, 1973). Like typical vertebrate melanosomes,
Pachymedusa melanosomes are also found in melanophores. However, 28
electron microscopically, Pachymedusa melanosomes appear to be huge and
"fuzzy" compared with the normal smooth, round structures. Further studies (Bagnara, Taylor and Prota, 1973; Bagnara and Ferris, 1974) revealed the Pachymedusa melanosome to be a compound organelle composed of an electron-dense, melanin-containing kernel surrounded by a halo of
"fuzzy" material. Light micrographs of unstained skin sections re vealed melanophores which were a deep red-brown in color. This unusual color was attributed to the "fuzzy" melanosomal material and chemical tests were begun to decipher the nature of the red-brown pigment.
Preliminary chemical examination of this pigment demonstrated its extreme insolubility in everything except dimethyl sulfoxide (DMSO) or alkali (0.1 N NaOH). The pigment, when extracted in DMSO is bright red, and remains so indefinitely. In sodium hydroxide, the extracted pigment is initially bright red but breaks down to an orange and finally a yellow compound upon prolonged exposure to alkaline condi tions (Bagnara et al., 1973). Initially, Bagnara (Bagnara and Ferris,
1974) coined the term rhodomelanochrome to describe this red pigment.
Additional work revealed that rhodomelanochrome was, structurally, a pteridine dimer, and absorption spectra together with organic chemical data led to the identification of rhodomelanochrome as pterorhodin
(Misuraca et al., 1977).
Rhodomelanochrome, while no longer tenable as a name for the red pigment, was nevertheless initially misidentified for several good reasons. First, the likelihood that the red pigment, which is found 29 in melanosomes, was a pteridine was not initially considered. After all, in nearly all vertebrates examined, pteridines are the main compo nents of pterinosomes in xanthophores, and the presence of either pterinosomes or other organelles such as reflecting platelets in melanophores, while known to occur, appears to be the exception rather than the rule (Bagnara, 1976). Thus, the red pigment was presumed to be melanin-related, and perhaps a phaeomelanin of some sort. This was further supported by the fact that leaf frog skins were fully capable of incorporating labelled DOPA and tyrosine, presumably into melanin
(Bagnara et al., 1973). However, once enough material was available for skin extractions, it became rapidly apparent that this pigment was in no way related to the melanins, but in fact fluoresced and absorbed ultraviolet light similar to the pteridines. By this time it was strongly suspected that the red pigment might in fact be pterorhodin, and comparison with authentic pterorhodin purchased from Tridom Chemical
Company fully supported these suspicions (Misuraca et al., 1977).
Remarkably, while pterorhodin had been originally identified as rhodopterin by Hopkins (1895) at the turn of the century, reports in the literature concerning pterorhodin since that time have been scarce, and its occurrence in nature is rather rare (Ziegler and Harmsen, 1969).
Until pterorhodin was identified in Pachymedusa skin, it had only been known to occur in a few insects, and then primarily as an eye pigment
(Pfleiderer, 1964). Thus, it had seemed highly unlikely that such a pigment could be the dominant pigment in the skins of adult leaf frogs.
Even more startling was the discovery of a morphologically similar melanosome and red pigment in other members of the sub-family 30
Phyllomedusinae, and in the Australian hylid, Litoria caerulea (Bagnara
and Ferris, 1975; Bagnara, Ferris and Taylor, 1976). Such findings are
believed to be important to the taxonomy and evolution of these frogs,
as well as having broad developmental and biochemical significance.
Developmental Biology of Pachymedusa
When these studies began very little was known about the
developmental biology of the Mexican leaf frog. One of the first
questions asked regarding this animal was whether or not the red pig
ment was present throughout its development. Surprisingly, when tad
poles were obtained and tested for the presence of pterorhodin by
immersing small pieces of skin in 0.1 N NaOH, all such tests were
negative. Closer examination revealed that pterorhodin first appears
in the skins of these animals at the very end of metamorphosis, when only a faint trace of tail remains. Once this tiny remnant disappears,
pterorhodin is detectable in copious amounts, leaching out of sodium hydroxide-treated skin in quantity.
From these results, it was initially believed that the appear ance of this pigment was probably hormonally controlled like many other events which occur during amphibian metamorphosis (Frieden and Just,
1970). From previous workers (Kollros and Kaltenbach, 1952) it was inferred that the local effects of thyroxine, the hormone responsible for amphibian metamorphosis, could be tested by implanting thyroxine- cholesterol pellets under the skin of tadpoles and checking at inter vals for the presence of pterorhodin. A series of such experiments were planned and carried out with results to be discussed later. 31
Since the developmental biology of pigmentation in this animal is the topic of this study, a more thorough discussion of amphibian development in general will be witheld until later. Suffice it to note at this point that initial suspicions concerning the hormonal control of pterorhodin deposition were entirely too simplistic. Instead, pterorhodin synthesis, and indeed pteridine synthesis in general, in
Pachymedusa presented a complex system consisting of both direct and indirect developmental cues.
Statement of the Problem
These introductory pages provide a detailed explanation of the rationale for the present study. It is the purpose of this study to present the developmental sequence of pteridine pigments in Pachymedusa dacnicolor, and in doing so, to infer certain specifics about the metabolism and developmental significance of these pigments, particu larly concerning the appearance of pterorhodin. Naturally, these pig ments were first identified. Then, since so little is known about the actual enzymatic synthesis of pteridines found in any frog, a superfi cial survey of these mechanisms was attempted here. Xanthine dehydro genase seemed the most likely candidate for some role in the pigmenta tion features of leaf frogs since it was already known to play a role in its excretory pattern. The availability of an inhibitor for XDH and its proposed metabolic position preceding the synthesis of ptero rhodin would allow, for the first time, confirmation of the actual pathway for pterorhodin biosynthesis. Finally, while hormonal studies proved to be somewhat inconclusive, they were responsible for more
intensive biochemical studies. The end result is a combination of fact
and speculation leading to some suggestions concerning the regulation of pteridine synthesis and pterorhodin deposition in Pachymedusa dacni- color. CHAPTER 2
MATERIALS AND METHODS
The sub-headings in this chapter reflect various aspects of the overall plan of study. The organization is such that specific proce dures are grouped according to particular aspects of experimental investigation.
Maintenance and Rearing of Animals
All of the research accomplished herein relied heavily upon the ability to breed and maintain Pachymedusa dacnicolor in the laboratory.
Since this animal had not previously been used in this capacity, and little was known of its growth requirements, details of the methods found to work best for maintaining these animals in this laboratory are described below.
Adults
Sexually mature specimens of Pachymedusa dacnicolor were either supplied by Southwestern Scientific Co., Tucson, Arizona, or collected in the field from the states of Sonora and Sinaloa, Mexico. Live adults were maintained as a breeding colony in two greenhouses in Tucson. Some frogs spent the winter months in an environmental chamber at the
University of Arizona. These animals were induced to breed "out-of season" by artificially manipulating day length, temperature and
33 34 humidity. The normal breeding season in the greenhouses begins in mid- to late June and extends through August.
Animals fed on live insects attracted to the greenhouses by ultraviolet lights. During the winter months, and in the case of animals maintained indoors, their diet consisted of commercially sup plied or laboratory-raised crickets.
Embryos and Larvae
All tadpoles used in these studies were obtained from spawnings from the adult breeding colony. Egg clutches were best maintained in incubators at 28°C. In general, clutches were removed from the green houses or environmental chamber soon after the completion of laying.
Maximum development of embryos was achieved when eggs were suspended over large pans of dechlorinated water in the incubators. This seemed to best simulate natural conditions in which eggs laid on leaves or trees over rain pools hatched into the water below. Embryos suspended above the water in incubators also hatched into the water below, usually after 4-6 days of development. Clutches placed Immediately in pans of water often exhibited considerable mortality. Development through hatching proceeded much more slowly in embryos grown in water, and often the development of entire clutches arrested sometime prior to hatching.
Newly hatched, swimming larvae were transferred to large out door tanks with a constantly flowing supply of charcoal-filtered water.
Tadpoles were fed ad libitum on washed, canned spinach. For some ex periments animals were reared in large plastic pans in 28°C incubators.
Water was changed every other day and animals were fed every day. 35
Metamorphosing animals (those with all four legs) were trans ferred to covered pans tilted at a 30° angle and containing a small amount of water. This accommodated the need that metamorphosing animals had to crawl out of the water and remain relatively dormant until the completion of the metamorphic process. When metamorphosis was complete, animals were maintained in a screen-enclosed cage with a water supply and fed on small crickets and Drosophila.
The Taylor-Kollros (1946) staging system for leopard frog larvae was applied to Pachymedusa tadpoles. Only minor modifications to accommodate tree frog peculiarities were necessary for this staging system to work.
Chemicals
The following compounds were purchased from Sigma: xanthine, hypoxanthine, NAD, phenazine methosulfate, nitro-blue tetrazolium, allopurinol, cellulose for thin layer chromatography (Sigmacell, Type
100), Dowex 1X8 (chloride)-400 mesh, Sephadex G-10 and G-25 (fine),
ECTEOLA-cellulose, phosphocellulose, xanthopterin, isoxanthopterin, biopterin, leucopterin, pterin, pterin-6-C00H, EDTA, ammonium sulfate and thyroxine. Acrylamide, BIS-acrylamide, TEMED, ammonium persulfate,
2-mercaptoethanol, and TRIS were purchased from Bio-Rad. Silica gel G
(Merck) was supplied by Scientific Manufacturing, Inc. (SMI).
For some experiments the allopurinol used was a generous gift from Dr. George Hitchings of the Burroughs-Wellcome Research Labs.
George Marshall (Chemistry Dept., University of Arizona) synthesized the 7-methyl-xanthopterin used in these studies. Synthesis was carried 36 out as outlined by Elion and Hitchings (1947). Authentic pterorhodin was purchased from Tridom Chemical Company, and d-erythro-neopterin was a gift from Dr. J. Matsumoto. MS-222 (tricaine methanesulfonate) was purchased from Crescent Research Chemicals, Inc., Scottsdale, Arizona.
All solvents, reagents and other chemicals were of commercial, analyti cal grade.
Experimental Procedures
A chronological presentation of experiments and experimental methods is outlined below. Preliminary experiments examining the local effects of thyroxine on Pachymedusa tadpole skins were accomplished.
This was followed by a developmental analysis of the pteridines in lar val and newly metamorphosed animals, and a detailed analysis and iden tification of the pteridines from adults. Finally, studies concerning the enzyme, xanthine dehydrogenase, were completed with emphasis on the possible role of XDH in the pigmentation of Pachymedusa.
Thyroxine Implants
Pellets consisting of a mixture of 10% thyroxine and cholesterol were kindly provided by Dr. Jane Kaltenbach. These pellets were used in an attempt to induce local metamorphosis in the dorsal skins of leaf frog tadpoles.
Groups of 10-20 animals (stages XII-XVI) were anesthetized by immersion in a weak solution of MS-222. A small puncture was made in the dorsal skin on one side of the animal using two pairs of fine watchmaker's forceps (no. 5). Then, a pellet was slipped into the 37
puncture and pushed a few mm up and under the skin, away from the wound.
Experimental animals received pellets containing 10% thyroxine-choles-
terol. One group of controls received cholesterol pellets alone, and
another group was anesthetized, punctured, but received no pellet.
Animals were reared at room temperature (22°C), and fresh water and food
were provided as needed (at least every other day).
Experimental animals were observed for outward signs of meta
morphosing skin, such as a change in the color or in the texture of the
skin over the pellet. All animals were terminated by over-anesthetizing
in MS-222 before reaching stage XXIII. At this time animals were
photographed and a small section of thyroxine-treated skin was immersed
in 0.1 N NaOH to test for the presence of pterorhodin. Another sample
of thyroxine-treated skin was fixed and given to Dr. Ferris for electron
microscopy. Skins from the untreated portion of experimental animals,
skins from those animals receiving only cholesterol pellets and skins
from untreated animals were also tested for pterorhodin and fixed for
electron microscopy. Samples from controls were taken at intervals
corresponding to those of thyroxine-treated animals.
Pteridine Extraction and Identification
An excellent review of pteridine extraction and identification
procedures is presented by Matsumoto, Bagnara and Taylor (1971).
Modifications of several of these procedures were used to extract
pteridines from leaf frog skin.
Ethanol Extracts. Dorsal skins of various stage Pachymedusa were dissected, pooled and weighed. All subsequent steps were carried 38 out in total darkness or reduced light to prevent photodegradation of labile pteridines. Skins were pre-treated either by heating in an 80°C water bath for 15-30 min or by stirring in a 10X (fresh weight) volume of 70% ethanol overnight. Next, those skins that were stirred overnight were transferred, ethanol included, to a Potter-Elvejhem glass homogen izing vessel fitted with a motor-driven teflon pestle. Grinding was accomplished mechanically at 1000 rpm. Heated skins were also ground in a 10X volume of 70% ethanol. Homogenization was continued until all of the skin remained in a fine suspension. The ethanol was then poured off into a centrifuge tube and another 10X volume of 70% ethanol was added to the remaining skin and rehomogenized. The extraction procedure was repeated a third and final time using a 5X volume of 70% ethanol. All of the ethanol extracts were pooled in capped centrifuge tubes and heated for an additional 30 min at 80°C.
Next, tubes were centrifuged at 1500 x g for 20 min. Super- natants were transferred to clean centrifuge tubes and treated with a
3X volume of chloroform. After centrifugation at 1000 x g for 10 min, the clear, epiphasic layer was removed to a tightly capped brown jar.
Mercaptoethanol was added to a final concentration of 0.01%. Chloro form treatment removed most of the ethanol, proteins, lipids and other impurities, leaving an extract containing pteridines, purines and other small molecular weight compounds.
In the case of large extracts (more than 0.25 g initial fresh tissue weight), fluorescence usually persisted in the chloroform layer after this step in the extraction procedure. When this occurred, a 39
small volume of distilled water (about 1 ml) was added to the remaining
chloroform. The tube was capped, thoroughly shaken and centrifuged
again at 1000 x g for 10 min. The clear, epiphasic layer was added to
the initial extract. In rare cases the chloroform was treated a second
time with water to remove the last traces of fluorescence. Fluorescence
was monitored visually by viewing extracts over an ultraviolet light
source equipped with a Corning (M2401) dark blue filter. Final pooled
extracts were stored frozen and in the dark.
Adult skin and skins from newly metamorphosed animals were
extracted by slightly modifying the above procedure to compensate for
the increased quantity and insolubility of the pteridines present.
Extraction was repeated a total of five times, twice using a 10X volume
of ethanol and three times using 5X volumes. When ethanol extraction was complete the remaining skin was re-extracted twice using a 5X vol
ume of 88% formic acid each time. Instead of homogenizing the already
finely ground skin, the formic acid was simply added, mixed well and
allowed to settle for 30 min. The bright red extract was centrifuged
at 1500 x g for 20 min and the formic acid extraction was repeated at
least one more. time. If red pigment continued to persist in the re
maining skin, formic acid extraction was continued until only a faint
pink color was observed. This harsh extraction procedure was necessary
in order to extract the extremely insoluble red pteridine dimer, ptero-
rhodin. Partial neutralization of formic acid was accomplished by
adding an equal volume of distilled water to the extract. Then the
entire extract was washed with an excess of anhydrous ethyl ether. 40
Pterorhodin can then be collected and purified as described by Misuraca et al. (1977).
Ammonium Hydroxide Extracts. Pteridine extraction was also accomplished using 0.5 M NH4OH in place of ethanol. In this case, stirring skins overnight in a 10X (fresh wt.) volume of 0.5 M NH4OH was sufficient to extract the majority of fluorescence even without homo- genization. Centrifugation was then carried out at high-speed (20,000 x g for 20 min) to remove dissolved melanin and non-fluorescent con taminants. To further ensure maximal extraction an additional 5X volume of NH^OH was used to wash the pellet after removal of the initial extract. The washing was added to the extract, and the entire prepara tion was treated with chloroform and centrifuged at 2,000 x g for 30 min. The epiphasic layer was treated as described above for the ethanol extract. Then, extracts were treated with a small volume of 10% tri chloracetic acid (TCA) to precipitate proteins. This preparation was centrifuged at 20,000 x g for 20 min, and the supernatant was saved.
In some cases extracts were lyophilized and resuspended in distilled water to a final concentration of 100 mg (initial tissue wt.)/ml.
Final extracts were stored frozen and in the dark.
This extraction procedure is not suitable for recovery of pterorhodin. Overnight exposure to alkaline conditions will result in considerable breakdown of the red pigment.
Thin Layer Chromatography (TLC). Pteridines are best visualized by subjecting them either to paper or thin layer chromatography. Since all of the extracts prepared from Pachymedusa skins were extremely 41 heterogeneous, some form of two-dimensional chromatography and/or electrophoresis had to be developed so that all ultraviolet fluorescing
(and absorbing) compounds could be clearly distinguished.
After a great deal of trial and error a suitable TLC media was developed which provided excellent one- or two-dimensional separation of all components from skin extracts. The significant difference in TLC as performed in these studies and standard TLC procedures was the prepara tion of plates from a 1:1 slurry of silica gel G and cellulose. This combination was found to produce excellent and reproducible separation of pteridines, and the rationale and details of the system are described in detail elsewhere (Frost and Bagnara, 1978).
For these studies chromatography was accomplished on 20 x 20 cm glass plates coated with the silica gel G:cellulose preparation. In some cases one-dimensional TLC was carried out on 5 x 20 cm plates. All chromatography was performed ascendingly in glass tanks with stainless steel plate holders purchased from SMI. In all cases the solvent used for chromatography in the first direction was n-propanol:7% ammonia
(2:1), and iso-propanol:2% ammonium acetate (2:1) for the second direction. Plates were always heat activated at 100°C for 10-20 min just prior to use. A Hamilton microliter syringe was used to apply 25-
30 ill of sample 1 cm from the bottom of the plate and 4 cm from the left edge in the case of two-dimensional TLC, or 1 cm from the bottom of the plate and at least 1 cm apart in the case of multiple samples and one- dimensional TLC. Chromatography was carried out until the solvent front had progressed at least 10 cm from the sample origin in both directions. 42
The entire two-dimensional procedure required less than four hours,
while one-dimensional TLC required about lh hours.
Results of chromatograms were evaluated by computing Rf values
for each spot in each direction. These values were then compared with
those from authentic pteridines exposed to the same chromatographic
conditions. This allowed the tentative identification of pteridines in
Pachymedusa skin extracts. A record of each chromatogram was kept by
photographing plates sitting on an ultraviolet, dark blue filtered light
source. A suitable photograph can be obtained using Kodachrome-25
film, and a 35 mm camera equipped with a 55 mm lens and No. 3 supple
mental lens. The camera should be set at f 1.8 with a one second
exposure. A diagrammatic representation constructed from the computed
Rf values was also kept as an additional record of TLC results.
Column Chromatography. An ethanol extract was prepared from
2.52 g (fresh wet weight) of adult Pachymedusa dacnicolor skin. The remaining skin residue was subjected to formic acid extraction. The
ethanol extract (about 50 ml) was lyophilized and resuspended in 15 ml of distilled water. This extract was then subjected to column chroma tography on a variety of ion exchangers as suggested by Rembold (1971).
First, the extract was run on a Dowex 1X8 (formate) column
(1 x 40 cm). Water was used as the elutant and 5 ml fractions (in the time mode) were collected using a Beckman Model 132 automatic fraction collector. When no further fluorescence was eluted with distilled water, 250 ml of 5% formic acid was washed through the column, followed by 250 ml of 10% formic acid. No additional fluorescence was eluted 43 with either stronger formic acid concentrations or formic acid/sodium formate solutions as suggested by Rembold (1971). All fractions were monitored for absorbance at 260 nm in a Beckman DU spectrophotometer and for fluorescence at 450 nm using a Beckman DU/DK fluorescence attachment on the spectrophotometer. Fluorescing fractions were also run one-dimensionally on thin layer plates as outlined in the previous section.
Water-eluted fractions from Dowex chromatography were treated in several ways. First, all similar ultraviolet absorbing and/or fluorescing fractions were pooled according to their contents as determined by TLC. Those pooled fractions less than 20 ml in volume were rechromatographed on a 1.5 x 45 cm column of phosphocellulose.
Pooled fractions greater than 20 ml in volume were lyophilized, resus- pended in a small volume of distilled water and rechromatographed either on the phosphocellulose column or on an ECTEOLA-cellulose (2.5 x 40 cm) column. Elution of these fractions was again accomplished using dis tilled water, and 5 ml aliquots were collected. Absorbing and fluor escing fractions were examined for purity both spectrophotometrically and on thin layer plates. These fractions were then pooled and lyophil ized. Pure fractions were resuspended in an appropriate solvent and an absorption spectrum was run as outlined below. Fractions with more than one substance were also lyophilized and resuspended in water.
However, these were then rechromatographed on a different column.
Formic acid fractions from the Dowex column were pooled accord ing to their contents and then extensively washed with ethyl ether to remove most of the formic acid. Pooled fractions were handled as de
scribed above except that rechromatography was always carried out on
ECTEOLA-cellulose, and in some cases a 1-5% gradient of formic acid was
utilized to elute the compounds from the column.
Once pure compounds were obtained and lyophilized, absorption
spectra were recorded in order for compounds to be positively identi
fied. Fluorescing compounds were dissolved either in 0.1 N NaOH or
0.1 N HC1. Absorption spectra were recorded from 210-440 nm in a
Beckman DB-G grating spectrophotometer equipped with a scanning feature and an automatic recorder. Spectra from extracted compounds were com pared with those from authentic compounds.
The remaining initial formic acid skin extract was not subjected to column chromatography, but instead was used to purify and identify pterorhodin. This procedure is outlined by Misuraca et al. (1977), and need not be further discussed. Purification of pterorhodin was followed by its identification from a comparison of absorption spectra of the compound isolated from Pachymedusa skins and of an authentic sample of red pigment.
Xanthine Dehydrogenase Experiments
The approaches utilized to assay and extract xanthine dehydro genase from Pachymedusa are very similar to those employed for Droso- phila (Seybold, 1974) and for chicken liver (Murison, 1969). Fortunate ly, xanthine dehydrogenase is a soluble enzyme, always of relatively high, molecular weight (usually around 300,000). Thus, most extraction procedures for this enzyme will not vary greatly from organism to 45 organism, and were found to work quite well in this study of the
Mexican leaf frog enzyme.
Crude Extraction. In a number of experiments only a crude tissue preparation was employed in the determination of xanthine dehy drogenase activity. All tissues used for extraction were dissected from freshly killed animals. Tadpoles were killed by immersing them in an ice bath. Adults and metamorphosed animals were killed by decapitation and pithing. Tissues were pooled until at least 0.4 g of each tissue
(usually liver, kidney and skin) was amassed. All subsequent procedures were carried out at 0-4°C.
Tissues were homogenized in a 10X (based on fresh wet weight) volume of 0.05 M TRIS-HC1, 0.001 M EDTA, pH 8 buffer (standard buffer).
Homogenization was accomplished using a Potter-Elvejhem glass homogen izing vessel fitted with a motor driven teflon pestle. Grinding was carried out at 1000 rpm for 30-60 sec (or longer if necessary), until a fine suspension was achieved. The homogenate was transferred to a centrifuge tube and centrifuged at 30,000 x g for 20 min at 4°C. The resulting supernatant was carefully drawn off the pellet and used directly for XDH assay.
Assays were usually carried out on the same day as extraction.
In some instances extracts were stored at 2°C and assayed 24-48 hours later. Storage at 2°C for as long as two weeks revealed no significant decline in enzyme activity.
In some cases serum was collected from animals and examined for enzyme activity. In this instance, freshly killed animals were bled directly from the heart into capillary tubes. Blood from the same stage 46
animals was pooled and diluted 1:1 with standard buffer. This prepara
tion was then centrifuged at high speed, and the resulting supernatant was assayed for enzyme activity.
Partial Purification. The first step in the partial purifica tion procedure was the preparation of a crude extract as described above. Then, a cold, saturated solution of ammonium sulfate in standard buffer was added to the extract to a final volume of 67% saturation.
This preparation was centrifuged at 30,000 x g for 20 min and the super natant was discarded. The resulting precipitate was redissolved in a
10X volume of standard buffer and ammonium sulfate precipitation was repeated. The final precipitate was dissolved in standard buffer to a final concentration of 100 mg (initial fresh weight)/ml. This extract was assayed for XDH activity as outlined below. Extracts were stored either at 2°C or frozen.
Enzyme Assay. The assay procedure for XDH (Murison, 1969) is based on the enzyme's ability to convert xanthine to uric acid. Uric acid absorbs ultraviolet light maximally at 290 nm (millimolar absorp tion coefficient for uric acid at 290 nm = 11), and its appearance can thus be monitored spectrophotometrically.
The assay solution consisted of 0.2 mM xanthine and 0.7 mM NAD in standard buffer. The reaction was initiated by adding 0.3 ml of extract to 3 ml of assay solution in a quartz cuvette. A blank was prepared by omitting xanthine from the above reaction mixture. The change in OD29Q was followed for 20 min in a Beckman DU spectrophoto meter. The reaction was found to be linear with time during this 47 period, and final enzyme activity was expressed in "units" (mumoles of uric acid produced/min/mg tissue fresh weight). All protein determina tions were done as outlined by Lowry et al. (1951).
Electrophoresis. Visualization of XDH was accomplished by sub jecting the enzyme to polyacrylamide gel electrophoresis and staining specifically for XDH (Yen and Glassman, 1965). Gel electrophoresis was carried out on a Bio-Rad Model 220 vertical slab gel apparatus equipped with a Buchler Model 3-1014A power supply. The buffer system and acryl- amide gel preparation were those suggested by Gabriel (1971). Gels were composed of 7.5% acrylamide and 0.2% BIS in TRIS-HC1 buffer, pH 8.6.
The electrode buffer consisted of 0.005 M TRIS, 0.03 M glycine, pH 8.8.
Samples (25-75 ul) were made 5% with respect to sucrose and applied to the sample slots with a microliter syringe. Electrophoresis was carried out for 16 hours at a constant voltage of 100 volts (about 12 mamps) at 2°C.
Gels were stained by incubating at room temperature in the fol lowing preparation: 1.0 mM hypoxanthine, 0.45 mM NAD, 0.26 mM phenazine methosulfate, and 0.39 mM nitro-blue tetrazolium in standard buffer.
Hypoxanthine was weighed out first and dissolved in a few drops of 1 N
NaOH. Buffer was added to this, followed by the other compounds, one at a time. The staining solution was made up just prior to use.
The enzyme began to appear on gels after about 30 minutes of staining at room temperature. Beyond 10 hours, staining was no longer effective, and the entire gel stained a uniform dark blue. Thus, after
6-10 hours, the reaction was terminated by pouring off the staining 48
solution and immersing the gel in 6% acetic acid. After 30 minutes,
the acid was poured off and the gel was washed several times with dis
tilled water. Gels were photographed and stored wrapped in plastic.
Inhibition. Several groups of experiments involving the inhibi
tion of XDH activity were accomplished using allopurinol. This drug was
administered to tadpoles by one of three methods. First, animals were
grown, beginning at 1-2 days post-hatching, in water containing 0.05%
allopurinol. Fresh water and drug were supplied every third day.
Secondly, animals (stages XVIII-XIX) were injected (i.p.) daily with
0.05 g/g body weight of allopurinol. The injection solution consisted
of allopurinol dissolved in boiling, dechlorinated tap water. Controls were injected with water alone. In a third set of experiments, animals
were fed a diet consisting of spinach, gelatin and allopurinol. This
special diet was prepared by heating approximately 200 ml of canned
spinach to boiling and gradually stirring in 1 g of gelatin and 1 g of allopurinol. The control diet was prepared in the same way except that
allopurinol was omitted. Animals maintained on this diet were fed at least once a day and fresh water was supplied every other day.
In all of the above experimental groups animals were reared to appropriate developmental stages and then sacrificed. Livers, kidneys and halves of each dorsal skin were pooled, extracted and assayed for
XDH activity. The remaining dorsal skin halves were pooled, extracted and analyzed for pteridine content. Animals, stage XXV or older were also tested for the presence of pterorhodin by immersing a small piece of skin in 0.1 N NaOH. Control animals were handled in the same manner. CHAPTER 3
RESULTS
The results are presented in an order which reflects the experi mental procedures already described and which lends to the coherence of the ensuing discussion. For convenience, most of the relevant data is presented in the form of figures, photographs and tables.
Thyroxine Experiments
Five separate groups of thyroxine implants were accomplished.
These experiments involved a total of 68 animals. Of these, 12 animals
(17.6%) failed to respond to the hormone, 19 (27.9%) responded to thy roxine by completing metamorphosis, and 37 (54.5%) demonstrated a local response to hormone as evidenced by the appearance of a "spot" in the vicinity of the pellet. None of the controls ever demonstrated any kind of a metamorphic response other than those naturally expected to occur with time. In general, a response to thyroxine occurred within
9-15 days post-pellet implantation. Animals failing to respond to thyroxine never exhibited a local "spot" and progressed developmentally at rates comparable to those of controls. Animals responding to hormone by completing metamorphosis exhibited a number of outward signs of precocious metamorphosis such as rapid emergence of somewhat under developed forelimbs, tail regression and the incomplete development of
49 adult mouth parts. All of these precociously transforming tadpoles died during or shortly after the completion of metamorphosis.
A spot typical of a local thyroxine response is shown in Figure
4. Such spots were initially distinguishable by their more intense color (green or red-brown) in contrast to either surrounding skin or skin of controls. Spots appeared to contain more pigment cells and skin glands than control skin of a comparable age and location. These were all outward indications of local metamorphosis. Animals exhibiting signs of local metamorphosis demonstrated few if any other signs of precocious metamorphosis.
All animals responding locally to thyroxine were sacrificed between stages XVIII and XXIV, (three stage XVIII, three stage XIX, eight stage XX, eight stage XXI, nine stage XXII, five stage XXIII and one stage XXIV). In all cases a portion of the skin was tested for the presence of pterorhodin. None of the animals tested for precocious pterorhodin synthesis demonstrated any positive results. Control skins younger than stage XXV also never demonstrated any evidence of the presence of pterorhodin, indicating that as far as precocious red pig ment synthesis was concerned, thyroxine had little or no direct effect upon the skin.
Pteridlne Analysis
Column chromatography of an adult skin extract led to the iden tification of pteridines from Pachymedusa dacnicolor• This information was sufficient to allow for subsequent identification of most of the pteridines extracted from the relatively small skin samples of various Figure 4. Local Effect of Thyroxine on Pachymedusa dacnicolor Skin.
This figure depicts a stage XX tadpole approximately eight days post- pellet implantation. Arrows point to the localized "spot". 52 developmental stages. The results of these pteridine analyses are summarized below.
Adult Pteridines
The lyophilized adult skin extract was resuspended in about 15 ml of distilled water and applied to a Dowex 1X8 (formate) column. The subsequent elution profile is illustrated in Figure 5. Strong peaks at
260 nm were generally attributable to purines or similar ultraviolet- absorbing compounds. Fluorescence at 450 nm was always attributable to pteridines. Often predominance of either a fluorescing or an absorbing compound obscured the overlap of both types of compounds in any single fraction. Thus, none of the fractions collected from the initial Dowex run were considered to be pure, and all were further purified by re- chromatography on a different ion exchanger, usually ECTEOLA-cellulose.
Results from two different rechromatography runs on cellulose columns are illustrated in Figure 6. In "A", Dowex fractions 190-220 were lyophilized, resuspended in 20 ml of distilled water and rechroma- tographed on an ECTEOLA-cellulose column, eluted with water. In "B", lyophilized Dowex fractions 30-48 were rechromatographed on a cellulose phosphate column. As can be seen, the elution profiles are somewhat similar. The elution pattern for a particular pteridine was dependent upon several factors, including the ion exchanger utilized for chroma tography, and the charge and molecular weight of the compound eluted.
In the water-eluted Dowex fractions, most of the pteridines were ini tially contaminated with a strong u.v. absorbing compound (later iden tified as guanine). During rechromatography on ECTEOLA or cellulose LigM yritow A94 AoO<«>aftg Rf • .90 7" CH,'XANTHOPTERIN
o n oa
ColorlfM Btua fluorescing R,« « PTERIN-(-CO OH
SO
Figure 5. Dowex 1X8 (Formate) Column Chromatography of an Adult Pachymedusa dacnlcolor Skin Extract.
An ethanol extract from adult Pachymedusa dacnlcolor skin was prepared as described In Chapter 2. This was layered on a Dowex column and elutlon was begun with water. The first 250 fractions were eluted with water, followed by 100 fractions eluted with 5% formic acid, and the remaining fractions with 10% formic acid. Subsequent identification of fluorescing compounds as described in the text is presented in this figure. Rf values from TLC as well as natural and fluorescing colors for each compound are also presented. 1.0 1.0
20
Figure 6. Column Chromatographic Purification of Two Pteridines from Pachymedusa dacnicolor Skin.
In A, Dowex fractions 190-220 were layered on an ECTEOLA-cellulose column and eluted with water. In B, Dowex fractions 30-48 were layered on a phosphocellulose column and eluted with water. The resulting profiles show the separation of u.v. absorbing ( ) from fluorescing ( ) compounds. A portion of each fluorescent peak was checked for purity by TLC as described in Chapter 2. 55
phosphate, the u.v. absorbing compound usually came off the column
ahead of any pteridines. In Figure 6A the separation of u.v. absorbing
material from fluorescing material was very clean, while in Figure 6B
there was some overlap. Before identification these contaminated frac
tions were rechromatographed a second time on phosphocellulose. This
time separation was complete and the pure fluorescing fractions were
combined and lyophilized. Lyophilized compounds were resuspended in
3 ml each of 0.1 N NaOH and/or 0.1 N HCl. In this way absorption spectra were recorded for each purified fluorescent compound, as illus trated in Figure 7 for the above-mentioned fractions (Fig. 6). For these two particular compounds the spectra obtained in NaOH were very similar. Final confirmation of the identity of these compounds as neopterin and pterin was assured only after a second spectrum in HCl was run and a comparison was made of the Rf values on TLC plates with those of the authentic compounds. Once identified, it was apparent that the size and charge difference between pterin and neopterin fully accounted for their different elution profiles, both on Dowex (Fig. 5) and on cellulose columns (Fig. 6).
In general, most pteridines do not have such similar spectral patterns, and a single absorption spectrum will suffice for identifica tion purposes. This particular example was utilized here to illustrate the variety of parameters necessary to positively identify a particular pteridine. In most cases purity was sufficient enough after one re- chromatographic procedure to identify the compound in question. In some instances fractions presumably containing the same pteridine were pooled 01 N HCI
MwfllK AMP ( ) Dowci frocltont 30-48 ( )
WAVELENGTH (ran) WAVELENGTH (mil)
Figure 7. Absorption Spectra of Two Purified Pteridines. ui ON and lyophilized prior to identification in order to accimulate enough of the substance for clear-cut identification. Still, for some compounds, only enough material for a single spectral pattern was obtained. Never theless, eight fluorescing compounds were identified from Pachymedusa skin: biopterin, neopterin, isoxanthopterin, xanthopterin, leucopterin, pterin, pterin-6-carboxylic acid and 7-methyl-xanthopterin. The posi tion of these pteridines as eluted on Dowex is shown back in Figure 5.
Traces of other fluorescent compounds (at least four) were not present in sufficient quantities to allow for their identification. The ninth pteridine, pterorhodin, was extracted directly from adult skin with 88% formic acid, partially purified as described in the last chapter, and identified by comparison of its absorption spectrum with that obtained from authentic pterorhodin. The resulting data, including absorption maxima, TLC Rf values and compound colors, used for the identification of these nine pteridines are presented in Table III. The same results for authentic compounds run simultaneously with and under the same conditions as extracted compounds are also presented in Table III for the sake of comparison.
Larval Pteridines
Once these nine pteridines were identified, the color and Rf values were in turn used to evaluate and identify thin layer chromato- grams of alkaline and ethanol skin extracts from various larval stage
Pachymedusa. Each extract was subjected to two-dimensional TLC. A representative chromatogram is illustrated in Figure 8. The photograph in this figure is that of an actual chromatogram. Diagrams were also 58
Table HI. Physico-Chemical Properties of the Pteridines Isolated and Purified from Pachymedusa dacnicolor Skin.
Natural Fluores. Rf Pterldine Color Color Absorption Spectra, max Value*
Leucopterln colorless blue 240,280,,340 (in 0.1 N NaOH) .08 (Authentic) colorless blue 240,282,,342 (in 0.1 N NaOH) .07
Biopterin colorless blue 252,360 (in 0.1 N NaOH) .51 262,322 (in 0.1 N HC1) (Authentic) colorless blue 254,363 (in 0.1 N NaOH) .51 263,232 (in 0.1 N HC1)
Keopterin colorless blue 254,365 (in 0.1 N NaOH) .40 247,322 (in 0.1 N HC1) (Authentic) colorless blue 255,365 (in 0.1 a NaOH) .38 248,322 (in 0.1 N HC1)
Pterin colorless blue' 254,362 (in 0.1 N NaOH) .50 245,313 (in 0.1 N HC1) (Authentic) colorless blue 253,363 (in 0.1 N NaOH) .50 243,313 (in 0.1 N HC1)
Pterin-6-C00H colorless blue 262,365 (in 0.1 N NaOH) .18 260,350 (in 0.1 N HC1) (Authentic) colorless blue 262,364 (in 0.1 N NaOH) .17 260,348 (in 0.1 N HC1)
Xanthopterin lemon yellow aqua 255,390 (in 0.1 N NaOH) .24 (Authentic) lemon yellow aqua 256,392 (in 0.1 N NaOH) .24
7-CH3~Xanthopterin lemon yellow aqua 255,345 (in 0.1 N NaOH) .30 (Authentic) lemon yellow aqua 253,344 (in 0.1 N NaOH) .31
Isoxanthopterin colorless violet 225,255, 340 (in 0.1 N NaOH) .27 288,340 (in 0.1 N HC1) (Authentic) colorless violet 225,255, 340 (in 0.1 N NaOH) .26 287,340 (in 0.1 N HC1)
Pterorhodin red deep blue 240,325,502,540 (in 88% formic acid) 0 (Authentic) red deep blue 238,324, 505,540 (in 88% formic acid) 0
*The solvent used for TLC was n-propanol:73! ammonia (2:1). 59
IXP**
Figure 8. Two-Dimensional Thin Layer Chromatography of an Early Larval Pachymedusa dacnicolor Skin Extract.
Chromatography was performed as described in Chapter 2. Individual pteridines are identified as follows: XP, xanthopterin; IXP, iso- xanthopterin; NP, neopterin; AHP, pterin; BP, biopterin. 60
constructed from the Rf values as calculated from the original chroma-
togram. In all cases a photographic and a diagrammatic record of
chromatograms was maintained as a double check on the end result. In
every experiment pteridines were evaluated qualitatively and quantita
tively. Qualitative judgement was based on color and Rf values, and in
general was sufficient to identify by name all predominant pteridines.
Quantitative evaluation was accomplished using an arbitrarily designated
scale of "+'s" to indicate relative quantity. Thus, a single "+" de
notes a pteridine present in barely detectable amounts, while "I I 1 I I"
indicates a predominant pteridine. Since all extracts in this study
were prepared in a similar manner, and since each chromatogram repre
sented 25-30 jil of extract, then the quantitative data recorded is
believed to be fairly representative. Also, with the exception of
animals younger than stage X, most of the stage specific extracts were
carried out a number of times and with similar results each time,
providing the extraction conditions were not varied. Figure 9 is a
photograph of a one-dimensional thin layer chromatogram depicting the
pteridine pattern of some representative developmental stages of leaf
frogs. Tables IV and V are detailed analyses of these patterns, in
cluding the qualitative and quantitative estimation of pteridines
present at each developmental stage. In this way a clear illustration
of the developmental variations in pteridines can be visualized.
The results of alkaline and ethanol extracts (Tables IV and V) are presented separately because there are some differences attributable solely to the extraction procedure employed. Regardless of how skins were extracted, it was apparent from these results that the most Figure 9. Thin Layer Chromatogram of Skin Pteridines from Representative Developmental Stages of Pachymedusa dacnlcolor.
From left to right the stages shown are: V, XI, XXI, XXII, XXIII, XXIV (bad sample), XXV, XXV (raised in allopurinol), and adult. Chromatography was performed as described in Chapter 2. 62
Table IV. Developmental Record of Alkaline Skin Extracts from Leaf Frogs
Stage of Ribo AHP-6- 7-CHo- Development flavin COOH XP IXP NP AHP BP XP LP Pt
I - ++ ++ -H-H- +4+ ++ 4-+ - -
XX - ++ ++ -H-H- 4-H- ++ -H- - -
III - ++ ++ -H-H- 4-H- ++ ++ - -
IV - ++ ++ +H- +++ ++ ++ - -
V - - -
VI - ++ ++ +++ +++ ++ ++ - -
VII - ++ ++ +++ +++ ++ +++ - -
VIII - 4-+ ++ +-H- +++ ++ +++ - -
IX - +++ +++ -H-H- +++ ++ ++ - -
X - +++ ++ ++++ +++ ++ ++ - -
XI - ++ ++ ++4+ 44-1- 4+ ++ - -
XII - ++ ++ •H-H- •H-h 4+ ++ - -
XIII - ++ ++ •H-H- +++ ++ ++ - -
XIV - +++ +++ •H-H- 4+++ ++ ++ - -
XV - +++ +4-f -H-H- 4++ ++ ++ - -
XVI - ++ ++ ++++ +++ ++ ++ - -
XVII - ++ ++ •H-H- +++ ++ ++ - -
XVIII - ++ ++ ++++ +++ ++ ++ - -
XIX - ++ -H- 4-H+ ++++ ++ ++ - -
XX - ++ +++ ++++ ++++ ++ ++ + -
XXI - + +++ +++++ ++++ ++ ++ + -
XXII - + -H-f +++++ ++++ ++ + + -
XXIII - ++ +++ ++44+ ++++ ++ + + -
XXIV - +++ +++ +4H-H- ++++ 4+ + ++ -
XXV - -H-+ +++ 44+4+ ++++ ++ + -H- + ++
XXV+ - 4-++ +++ ++++ +++ ++ + ++ ++ 1 1 1 1 1
Adult _ +++ -H-H- ++++ 4-H- + +++ ++++ ++ 1 1 1 1 1
Relative quantities of each pterldlnes are designated by +'s. 63
Table V. Developmental Record of Ethanol Skin Extracts from Leaf Frogs
Stage of Ribo AHP-6- 7-CH3- Development flavin COOH XP IXP NP AHP BP XP LP PC .
I ++ ++ 4-f MM 4++ 4+4+ ++ - -
II ++ 4-f 4-f 1 1 1 1 +++ +44+ 4+ - -
III ++ ++ 4-f MM +++ 4+4+ 4+ - -
IV ++ 4-f 4-f 4-f4» +++ 4+ 4+ - -
V 4-H- -H» 4-f 4-H- +++ 4+ 4+ - -
VI ++ ++ 4-f 4-H- 4+4+ 4+ 4+ - -
VII ++ 4-f 4-H- 4+4+ +4+ 4+ - -
VIII ++ + 4-f 4-H- 4+4-f 4++ 4+ - -
IX ++ + 4-f M M 4-H+ 4++ 4-f - -
X ++ 4-f 4-f MM 4+4+ +4+ 4+ - -
XI ++ 4-f 4-f 4-H-f 4++ 4+ 4+ - -
XII ++ 4-f 4-f "H-H-f 4++ 4++ 4+ - -
XIII ++ ++ ++ +++++ 4+4+ 4++ 4+ - -
XIV ++ 4-f 4-f +++++ 4+4+ 4++ 4+ - -
XV -H- 4-f +4- ++4++ 4+4+ 4+ 4-f - -
XVI ++ 4-f ++ -t-H-H- 4+4+ 4+ 4+ - -
XVII -W- 4-f 4-f 4+4++ 4+4+ 4++ 4+ - -
XVIII +• 4-f 4-f 4-H-f 4+4+ 4+ 4+ - -
XIX + + ++ 4+4+ 4+4-f 4-f 4+ - -
XX + 4-f 4-f 4+4+ 4+4+ 4+ 4+ + -
XXI + ++ -H- 4+4+ 1 II 1 4+ 4+ + -
XXII + + 4-f 4-H+ 4+++ 4+ 4-f + -
XXIII + + 4-4-f 4+4+ 4+4+ 4-f 4+ + -
XXIV + + +++ 4+4+ 4+++ 4+ 4-f 4-f -
XXV - 4- 4-H- 4++4-f 4+4+ 4-f + 4-f + 4+
XXV+ - + 4-H- +f+++ 4+4+ 4+ + +4+ 4-f 1 1 1 1 1
Adult - 4-f 4-H- 4+4++ 4+4-f + 4++ 4++ 4-f 1 1 1 1 1 1 1 1 1 1
RelaClve quanclcies of each pceridlne are deslgnaced by + 's. striking pteridine change during Pachymedusa development was the sudden appearance of 7-methyl-xanthopterin at stage XX, but never before this time. Isoxanthopterin and neopterin are the most prominent of the lar val skin pteridines and there are some quantitative fluctuations in these two compounds throughout development. In ethanol extracts there are considerable fluctuations in the quantity of pterin present from stage to stage. In alkaline extracts there is noticeably less fluctua tion in pterin but more so in the stage-to-stage evaluation of pterin-6 carboxylic acid. This probably reflects more degradation of pterin as result of the alkaline extraction procedure, rather than actual synthe sis of pterin-6-carboxylic acid. Also, riboflavin is only visible in
70% ethanol extracts and is never found in alkaline extracts. The reason for this is not clear but may reflect the degradation of ribo flavin in alkali. Ethanol extracts clearly show that riboflavin is present in significant quantity in young larvae, but fades with age until it disappears entirely by the end of metamorphosis. Leucopterin appears faintly at stage XXV, regardless of the extraction procedure employed, and is never found in very significant quantities even after the completion of metamorphosis. Again, leucopterin may be significant only as a natural breakdown product of pterorhodin, rather than as a natural precursor. This possibility and others will be discussed in detail in the next chapter.
Pterorhodin is not detected in skins either by thin layer chromatography or by the sodium hydroxide test prior to stage XXV.
Subsequent to this time pterorhodin rapidly increases quantitatively until it is clearly the predominant pteridine in adult skin. 65
Finally, it should be noted that xanthopterin increases quan titatively at stage XX in alkaline extracts and at stage XXIII in ethanol extracts. Also, while there are developmental fluctuations in biopterin, the significance of these in relation to the occurrence of other leaf frog pteridines is unclear. The developmental differences, those differences inherent to the extraction procedures employed and the reasons for these differences, will be discussed in the next chapter.
Xanthine Dehydrogenase Studies
Experiments were carried out first to determine whether leaf frog XDH could be extracted and assayed. Then, the tissue localization and developmental variations in XDH activity were examined. Finally, an attempt was made to determine what, if any role XDH plays in the synthesis of Pachymedusa pteridines.
Extraction, Characterization and Assay of XDH
Following the procedures outlined in the last chapter for the assay and extraction of XDH from various tissues, considerable amounts of enzyme were always detected in the livers of Pachymedusa dacnicolor throughout development. Less activity appeared to be present in kidney and still less (often none) in the skin. With this in mind, two pre liminary experiments were performed, one to determine whether any tissue specific XDH isozymes exist, and one to determine whether any or all of these enzymes are capable of converting pteridines.
Figure 10 is an electropherogram of enzyme from liver, kidney, serum and skin extracts. The conditions for electrophoresis and enzyme 66
Figure 10. Electrophoretic Analysis of Xanthine Dehydrogenase from Various Tissues of Pachymedusa dacnicolor.
Electrophoresis and staining specific for xanthine dehydrogenase are described in Chapter 2. From left to right the particular enzyme extracts employed came from: adult liver, stage XVIII liver, stage XXIII serum, stage XV kidney, and stage XXV skin. visualization were described in Chapter 2. In every case examined, and
regardless of the developmental stage, all enzyme activity was localized
in a single, distinct band. Moreover, in all tissues the migration of
this band was always the same. Figure 11 is a thin layer chromatogram demonstrating the ability of liver XDH (and other tissue XDH's) to convert pterin to isoxanthopterin. Partially purified extracts were incubated in standard buffer containing 10~% pterin and 10~^ M NAD.
If either of these components was lacking, the reaction failed to pro ceed. Pterin alone revealed one rapidly migrating, blue fluorescent spot on a thin layer chromatogram. Unfortunately, at neutral pH
(as in Fig. 11) the blue fluorescence characteristic of pterin is virtually invisible. Together, extract, NAD and pterin produced a slower migrating, purple fluorescent spot on thin layer plates in addi tion to the faint blue pterin spot. This purple spot was indistin guishable from authentic isoxanthopterin.
For the majority of the enzyme experiments the partial purifi cation procedure was employed for the determination of enzyme activities.
Repeated failure to detect activity in skins led to the use of simple crude extracts for enzyme determinations, and these were found in general to yield considerably more detectable enzyme than the partially purified extracts. Finally, crude extracts were utilized in an attempt to determine enzyme in sera from tadpoles. These results, while still in a preliminary state, are extremely significant to this study, justifying their rather premature presentation here. Figure 11. Pterldine Conversion by Pachymedusa dacnicolor Xanthine Dehydrogenase.
This figure represents a thin layer chromatogram of five enzyme extracts with and without the addition of pterin and HAD. The five extracts employed are the same as those shown in Fig. 10. A single fluorescing spot was produced In each case. These spots were identical with authentic isoxanthopterin (IXP). Blank spaces between each spot are where control samples without pterin were run. Unfor tunately, pterin alone does not fluoresce at neutral pH and thus was not seen on this chromatogram. 69
Figure 12 illustrates the fluctuations observed in the liver enzyme during development. The data in this figure were obtained solely from partially purified extracts, and the "error bars" reflect the cal culated standard deviation from the mean. While the overlap from one developmental stage to the next is considerable, the general trend in the liver enzyme throughout development is undeniably clear. Each point in Figure 12 represents results from 2-7 experimental groups with each group consisting of at least three and as many as 24 animals.
Figure 13 represents similar enzyme data from kidneys and skins. While these results are certainly questionable on the basis of the wide standard deviations, the trends are clearly suggestive and important to the present study. This importance will be considered in depth in the next chapter.
Finally, preliminary results of crude extracts from serum and skin of metamorphosing animals are presented in Table VI. Only data from metamorphosing animals is presented here for two reasons: (1) At least 14 "fair-sized" animals were needed to obtain enough serum for assay and analysis, and (2) the significant observations, at least of skin activity, occur primarily during metamorphosis. To date the accum ulation of enough of the same stage animals younger than stage XVI all at one time has been somewhat impractical. However, it is anticipated that with continued successful breeding, similar data from at least representative stages will eventually be accumulated. For now, the data presented in Table VI suggest a dramatic increase in serum enzyme levels around stage XXIII, as well as a gradual but steady rise in skin 70
.33-
.30
.03-
V X XV XX XXV adult STAGE OF DEVELOPMENT
Figure 12. Developmental Fluctuations in Leaf Frog Liver XDH. .25
.20
.05- 72
Table VI. XDH Activity from the Skin and Serum of Metamorphosing Pachymedusa dacnicolor as Determined from Crude Tissue Extracts.
Stage of Development No. Animals Units of XDH Activity
Skin Serum
XIX 22 .005 _*
XX 10 .002 _*
XXI 15 .007 .016
XXII 14 .007 .042
XXIII 15 .011 .135
XXIV 21 .005 _*
XXV 22 .018 .018
XXV+ 26 .019 .017
*No serum was collected for this stage animal. 73
enzyme levels during and beyond the climax of metamorphosis. These con
sistent but low levels of enzyme were previously undetected by the
partial purification procedure. The significance of these findings will
be discussed in the next chapter.
Inhibition of XDH
Several types of experiments utilizing allopurinol as an inhi
bitor of xanthine dehydrogenase were attempted during the course of this
study. Results from these enzyme inhibition experiments are presented
in the form of two tables (Tables VII and VIII). In the first of these
tables, the experiments performed involved either the injection of
allopurinol into larvae or their maintenance throughout larval life in
water containing allopurinol. It is clear from the results of all of
these experiments involving partially purified enzyme extracts that inhibition does occur, particularly in the liver and kidney. Again,
results from skin were somewhat ambiguous, and crude extracts were
employed to alleviate this problem. Table VIII presents the results of crude enzyme extraction data from allopurinol-treated animals. In this case, animals were maintained on a steady diet of allopurinol from the
time of their initial feeding until their death. Once again, the results are striking, with substantial decreases in enzyme activity in every case.
Finally, the effect of allopurinol on specific pteridines was examined by thin layer chromatography of ethanol skin extracts from control and experimental animals. Figure 14 is an example of a one- dimensional TLC comparison of early larval control vs. allopurinol 74
Table VII. The Inhibition of Xanthine Dehydrogenase Activity During Pachymedusa dacnicolor Development as Determined by Partially Purified Enzyme Extracts.
No. of Stage of Development Animals Units of XDH Activity*
Liver Kidney Skin
Early Larval AP** 10 .005 0 0
Early Larval Control 10 .038 .011 0
Mid-Larval AP 4 0 .007 .005
Mid-Larval Control 4 .135 .023 .004
Stage XXV AP inj. 7 .004 0 0
Stage XXV AP 3 .002 0 .005
Stage XXV Control 3 .174 .079 0
Stage XXV+ AP inj. 6 .011 0 0
Stage XXV+ AP 4 .006 0 0
Stage XXV+ Control 4 .138 .090 0
*Units of enzyme are expressed as numioles of uric acid produced/ min/mg (tissue wet weight).
**Experimental animals were either grown in a solution of allo- purinol (AP), or injected with allopurinol (AP inj.). See Chapter 2 for specific experimental protocol. 75
Table VIII. The Inhibition of Xanthine Dehydrogenase Activity During Pachymedusa dacnicolor Development as Determined by Crude Enzyme Extracts.
No. of Stage of Development Animals Units of XDH Activity*
Liver Kidney Skin
Early Larval AP** 7 .020 0 0
Early Larval Control 7 .129 .005 0
Mid-Larval AP 4 .017 0 0
Mid-Larval Control 4 .244 .069 0
Stage XXV AP 4 .149 .020 0
Stage XXV Control 4 .269 .122 .018
Stage XXV+ AP 4 .087 .009 0
Stage XXV+ Control 4 .220 .092 .019
*Units of enzyme are expressed as mumoles of uric acid produced/ min/mg (tissue wet weight).
**Experimental animals (AP) were fed a diet consisting of allo- purinol, spinach and gelatin. See Chapter 2 for specific experimental protocol. 76
W M
¥ »
Figure 14. Thin Layer Chromatographic Comparison of Early Larval Control vs. Allopurinol-Treated Pachymedusa dacnicolor Skin Extracts.
Control skin Is represented on the left and allopurinol-treated skin on the right. Note the increased pterin (AHP), the decreased iso- xanthopterln (IXP), and the total absence of the slowest migrating pteridine in allopurinol-treated skin. treated animals. These particular animals had been grown in a solution of allopurinol. Figure 15 is a composite of two-dimensional chromato- grams illustrating the differences between control and drug-treated animals at representative developmental stages (early larval, mid-larval and stage XXV-XXV+). Again, these animals had all been reared in a solution of allopurinol.
Of particular note in all of these chromatograms are the two pteridines, isoxanthopterin and pterin. As expected, isoxanthopterin, which is usually a predominant pteridine, was generally quantitatively reduced in drug-treated animals, while pterin was quantitatively in creased. Slight decreases in xanthopterin were also noted as well as the complete disappearance of an unidentified pteridine (possibly 7,8- dihydroxanthopterin) in allopurinol-treated animals.
All animals reaching stage XXV or XXV+ were tested for ptero- rhodin. In 72% of those animals which had been exposed to allopurinol throughout their larval life, the appearance of pterorhodin was re tarded generally from 1-3 days beyond the normal time of appearance of red pigment. Better or complete results were not obtained for several reasons. First, animals characteristically crawl out of the water and stop feeding by stage XX or XXI, in order to complete the metamorphic process. No effective means of administering allopurinol to animals beyond stage XX was found. Injections at this point in development were generally lethal to the already naturally stressed animals. Had inhibition been continued, then perhaps an indefinitely prolonged in hibition of pterorhodin appearance might have been achieved. That 72% 78
t
Figure 15. Alterations between Control and Allopurinol- Treated Skins during Pachymedusa dacnicolor Development.
Six separate two-dimensional chromatograms are shown above. The three on the left represent the effects of allopurinol treatment on Pachymedusa pteridines while the three on the right are from normal control animals. From bottom to top, the lower two chromatograms are from early larval animals, the middle two from mid-larval animals, and the top two from stage XXV-XXV+ animals. Note the general de crease in all pteridines in allopurinol-treated animals. Arrows begin at the origin and indicate the direction of migration. 79 successful inhibition was achieved in these experiments is certainly more than expected since the experiments which served as a basis for those reported here were only 50% successful. Such experiments involved growing Drosophila on media containing .05-.08% allopurinol and examin ing the resulting adults for outward characteristics of the rosy mutant
(Boni et al., 1967). Their best results only produced 50% phenocopies.
The effect of allopurinol on the other pteridines present at stage XXV-XXV+ in leaf frogs can be seen in Figure 15. While marked decreases in isoxanthopterin, xanthopterin and 7-methyl-xanthopterin are noticeable, again, these might have been more prominent had inhi bition been continued throughout metamorphosis.
As a final note to these inhibitory studies, it should be men tioned that within the limitations of these experiments, it appears that allopurinol simply inhibits the activity of XDH. There is no evidence at present to indicate that the actual synthesis of XDH is inhibited or altered in any way. CHAPTER 4
DISCUSSION
In order to facilitate the organization and comprehension of the discussion this chapter is divided into four sections. The first three sections concern theories and speculation arising from the experimental results presented in the last chapter. The final section summarizes the information derived from this study.
Thyroxine and Pterorhodin Deposition
The results obtained with thyroxine implants were initially disappointing and somewhat puzzling. The natural appearance of ptero rhodin at the very end of metamorphic climax seemed to be an indication of a close connection between red pigment deposition and hormonally controlled metamorphosis. The fact that local thyroxine implants failed in every case to induce precocious pterorhodin synthesis led to some interesting speculation.
That skin was capable of responding to hormone was clearly demonstrated by the appearance of "spots" exhibiting increased numbers of skin glands and pigment cells. Moreover, some animals responded to thyroxine by completing metamorphosis. Of the 19 animals responding in this way, none lived long enough to be tested for pterorhodin at the time of its natural appearance (stage XXV). All of these animals were
80 tested for pterorhodin deposition at the time of their death, with
entirely negative results.
From the literature, several significant facts came to light in
relation to the seemingly discouraging results obtained from the local
metamorphosis experiments. In the first place, the implanted pellets, while able to elicit a visible effect, may not have been sufficient to raise the threshold of thyroxine to the level necessary for pterorhodin synthesis. This possibility rests on the theory that during metamor
phosis, different tissues respond at different times and to different levels of circulating thyroid hormones, an idea first suggested by
Kollros (1961). Secondly, not all metamorphic events respond to the artificial administration of hormone as they would naturally (Little and Castro, 1976). A good example of this is the amino acid deriva tive, taurine. Like pteridines, taurine is a small molecular weight compound. It arises as an oxidation product of cysteine metabolism.
Taurine occurs naturally as a component of bile salts in conjugation with steroids, and it is believed to play some role in the etiology of muscular dystrophy. In bullfrog tadpoles taurine levels exhibit a marked increase during spontaneous metamorphosis, but this increase is eliminated if metamorphosis is artificially induced with triiodothy ronine. The authors point out that many, but not all biochemical, metamorphic changes can be induced in premetamorphie animals by thyroid hormone administration. They caution against investigation of in vitro or artificial metamorphic events without having first ascertained their natural occurrence. 82
So, the role of thyroxine in the direct deposition of ptero-
rhodin has not been entirely eliminated. Preliminary results from
several experiments currently being conducted in this laboratory lend
support to the contention that thyroxine is somehow involved in ptero-
rhodin synthesis. A single hypophysectomized Pachymedusa tadpole,
hormonally induced to metamorphose, demonstrated "normal" appearance
of pterorhodin at stage XXV. Organ cultures of larval Pachymedusa
skin were attempted in anticipation of artificially inducing ptero
rhodin synthesis in vitro. Results from such experiments revealed that
skins from stage XX or older animals would eventually synthesize low
levels of pterorhodin on their own if left in culture long enough. The
older the isolated skin, the more rapidly pterorhodin will appear.
Prior to stage XX, pterorhodin has not been found to ever appear in
cultures, and this includes cultures which have been treated with
thyroxine and/or possible pteridine precursors (Hiroyuki Ide, personal
communication, Research Assoc., Dept. of Cell. & Dev. Biol., U. Arizona).
Thus, a more likely role for thyroxine in the synthesis of red
pigment might be an indirect one. Such an indirect role might conceiv
ably stem from several sources: (1) from some factor outside of the
skin itself; (2) at some step preceding the actual synthesis of ptero rhodin but still within the skin; or (3) at a step preceding ptero rhodin synthesis but apart from the skin itself. The results from
organ culture experiments seem to have eliminated "2" as a possibility.
In the following pages, the investigation of pigmentary and biochemical
events associated with Pachymedusa development will be discussed in 83
relation to the question of which of the above remaining suggestions
might occur in the Mexican leaf frog.
Pterorhodin, Pteridine Precursors and Leaf Frog Development
If one examines the literature concerning pteridines, it rapidly
becomes apparent that little is known about the actual in vivo synthesis of pterorhodin in any organism. Two specific points concerning ptero rhodin synthesis must be addressed in this discussion. The first point concerns the chemical synthesis of pterorhodin as reported in the literature, and the second concerns the natural synthesis of pterorhodin as speculated upon in the literature and in the present study.
The implication from the literature is that pterorhodin is the terminal product of C-7 pteridine metabolism. Chemically, pterorhodin is readily synthesized from xanthopterin and 7-methyl-xanthopterin in a test tube (Russell et al., 1949). Unfortunately, this condensation occurs only in hydrochloric acid at a pH of 2-3. Thus, either an enzymatic mechanism exists for the intracellular synthesis of ptero rhodin, or the organellar depositories of pterorhodin (i.e., melano- somes) satisfy the low pH requirement. Either of these possibilities is likely and, to date, neither can be eliminated.
Another problem which stems from the literature is the question of naturally occurring 7-methyl-xanthopterin (chrysopterin). This com pound is believed not to occur naturally, but instead to arise via the hydrolytic breakdown of erythropterin. This belief is based in part upon the finding that whenever 7-methyl-xanthopterin has been found to occur, so has erythropterin (Pfleiderer, 1964). However, in Pachymedusa 84
7-methyl-xanthopterin would appear to occur naturally in the absence of
erythropterin, since erythropterin has not been found in any leaf frog
skin extract. Moreover, 7-methyl-xanthopterin first appears at stage
XX, a developmentally significant point during anuran metamorphosis in
general.
In most anurans, the abrupt biochemical and morphological
changes commonly associated with thyroid hormone response occur between
stages XX and XXIII (Frieden and Just, 1970). Such changes are paral leled by rather abrupt increases in circulating levels of thyroid hor mone (Just, 1968). Thus, in Pachymedusa, while pterorhodin appears somewhat later in development than expected, its precursor, 7-methyl- xanthopterin, appears and increases in a fashion strikingly similar to other hormone-mediated metamorphic events.
As noted previously, the unnaturally low pH required for spon taneous "test-tube" formation of pterorhodin from its precursors may be an indication of an as yet unidentified enzymatic mechanism in cells responsible for natural pterorhodin synthesis. Adult leaf frog skin would certainly seem to be an ideal system for examining the enzymatic requirements for pterorhodin synthesis. Moreover, two facts uncovered in these studies indicate that such an enzyme might be the limiting factor in pterorhodin synthesis. First, 7-methyl-xanthopterin appears at stage XX, at least 3-7 days prior to the first indications of pterorhodin (stage XXV). Xanthopterin, the other purported precursor is present throughout development, eliminating the lack of precursor compounds as a likely limiting factor. Secondly, pterorhodin occurs very abruptly at stage XXV, with such rapid subsequent quantitative increases that substantial differences in the amount of pterorhodin present in skin are detectable literally only hours after the complete disappearance of the tail remnant (stage XXV+). However, the evidence in favor of the final step in pterorhodin synthesis as the limiting step is still entirely circumstantial. Alternatives to this possibil ity are presented below.
It should be noted that there may be more than one step limiting pterorhodin synthesis, although other points of regulation must neces sarily precede the direct synthesis of red pigment. The possibility of an enzyme mechanism existing for the production of 7-methyl-xanthopterin from xanthopterin or some other related compound must also be consi dered. Such an enzyme might be closely regulated by thyroxine and the onset of metamorphosis, and thus, could be an excellent candidate for further developmental and regulatory studies.
Again, it is still very possible that once pteridine precursors are made available to the melanosome, pterorhodin deposition may occur by spontaneous condensation. Then, only the availability of precursors and their subsequent passage into the melanosome remain as limiting factors. To some extent, available precursors have already been elimi nated as limiting factors. Thus, the possibility of some genetically controlled transport phenomenon should be considered. It is certainly possible that some mechanism, perhaps similar to the protein receptors responsible for intracellular steroid hormone transport, may exist for the transport of pteridines from their site of synthesis to melanosomes. 86
Of course, this assumes that pteridines are not synthesized but may eventually be deposited within melanosomes. The validity of such sug gestions remains to be tested.
The developmental pteridine data presented in the last chapter suggest an alternative route for pterorhodin synthesis. Such a pathway might involve the pteridines leucopterin and 7-methyl-xanthopterin or leucopterin and xanthopterin. Those pteridines which might participate in the synthesis of pterorhodin are illustrated in Figure 16.
Leucopterin, like chrysopterin and pterorhodin, is a pteridine common to invertebrates, and until now, none of these compounds have been reported to occur in vertebrates (Ziegler and Harmsen, 1969).
Leucopterin is known to be synthesized enzymatically by xanthine dehy drogenase (Glassman, 1965a). In Pachymedusa traces of leucopterin appear at stage XXV; slightly more is present by stage XXV+, and even greater quantities occur in the adult. Its appearance always parallels that of pterorhodin. Unfortunately, it is difficult to determine whether leucopterin plays any role in the synthesis of pterorhodin since it is also one of the primary products of alkaline degradation of ptero rhodin. Natural oxidative decomposition of pterorhodin during extrac tion might be just enough to account for all of the leucopterin found at these developmental stages.
Several significant problems are suggested at this point. First of all, since xanthopterin is present throughout development as the pteridine analyses have revealed, and if xanthine dehydrogenase is present throughout development as these studies suggest, then it would ^OH
N' 7-CHj-xonlhopl«rln xanthopterin
sy 3 CHrC-COOH N N
7-CH3-xanthopterin J erythropterin
PTERORHODIN SN- leucopterin leucopterin
"NX OH
CH*C-COOH N N erythropterin xanthopterin
Figure 16. Pteridines Believed to be Involved in the Synthesis of Pterorhodin. 88 seem unlikely for XDH to fail to convert xanthopterin to leucopterin at any available opportunity. This is particularly true since xanthopterin is believed to be located intracellularly with other unconjugated pteri- dines. Also, these animals always contain significant quantities of isoxanthopterin, another pteridine reliant upon XDH for its formation.
However, if xanthopterin were compartmentalized, perhaps in melanosomes and possibly to be used later as a precursor for pterorhodin synthesis, then XDH might not be able to attack this particular pteridine. Cer tainly this as well as other possibilities deserve further considera tion.
Very little is known about the turnover or stability of pteri- dines, or of their intracellular synthesis and deposition. Several possibilities regarding this point are suggested by experiments from this study.
The results of experiments involving the inhibition of XDH and ensuing effects on pteridine synthesis may answer some questions con cerning pteridine turnover. In all inhibition experiments, drug ad ministration was always initiated shortly after hatching. By this time pigment cells had already migrated from the neural crest and were visi bly differentiated in the swimming larvae. Superficial examination re vealed a variety of pteridines already present by this time. Analysis of XDH-sensitive pteridines (i.e., isoxanthopterin, and to some extent, xanthopterin) revealed decreases in the quantities of these compounds when drug-treated animals were compared to controls. However, there was always a substantial amount of these pteridines present in skin despite the inhibition of the enzyme responsible for their synthesis. Two
plausible explanations can be offered for this: (1) Pteridine turnover
might be so slow and these compounds may be so stable that overall syn
thesis during development is negligible except for what may be needed
to maintain general body growth and subsequent multiplication of exist
ing pigment cells. (2) Allopurinol inhibition of XDH is competitive.
Moreover, all methods of administration attempted to date do not ensure
a constant dose level to all animals at all times. Thus, inhibition in
a particular animals may be "leaky", resulting in continued but de
creased synthesis of XDH-sensitive pteridines. Also, both of these
possibilities may exist to some extent, and further investigation will
be necessary in order to establish which of them exists in the natural state.
In insects and other invertebrates, organs such as the eyes, cuticle and wings are major storehouses of pteridines. While these organs may or may not be the actual sites of synthesis of pteridines, other organs such as Malpighian tubules, testes and fat bodies are known to synthesize pteridines, and for the most part, to excrete them into the haemolymph or out of the insect's body. Thus, in insects pteridines are believed to be excretory products with the additional function of serving as pigments. Their excretory role stems both from their ability to be excreted as waste products and from their extremely insoluble nature such that, if they are not excreted then they must be deposited somewhere to avoid the problems of toxicity and subsequent lethality (Harmsen, 1966). 90
In vertebrates, pteridines might play similar roles, particular
ly since in humans they were first discovered as components of human
urine (Fukushima and Shiota, 1973). Presumably, all vertebrates excrete
some pteridines, the level and kinds of pteridines being characteristic
for a particular organism in a particular state of health (Halpern et
al., 1977).
In lower vertebrates employing pteridines as pigments, such
compounds are generally localized within a particular layer of the skin
located between the stratum spongiosum and the stratum compactum (Hama
and Obika, 1959). Such a layer may be likened to a "graveyard" or depository for fluorescing compounds. Also, lower vertebrates (fish, amphibians and reptiles) are known to sequester pteridines in specific pigment cells of the skin (Bagnara and Hadley, 1973).
In leaf frogs, pterorhodin (a pteridine) is localized within melanosomes. Pteridines in general are believed to reside within pterinosomes. The enzymes necessary for the synthesis of unconjugated pteridines (see Fig. 2 and Table I), including xanthine dehydrogenase, are all soluble cytoplasmic proteins. Thus, while a xanthophore may contain the enzymatic mechanisms for synthesizing pteridines it must also contain a mechanism for transporting pteridines and for compart mentalizing them within their respective organelle, the pterinosome.
Whether melanophores can also synthesize and transport pteridines is purely a matter of speculation.
During the course of these studies, it was discovered that while xanthopterin, 7-methyl-xanthopterin and leucopterin were readily and 91 consistently extracted from skins with ammonium hydroxide, no traces of these C-7 pteridines were found if skins were extracted with ethanol at room temperature. Unless ethanol extracts were subjected to gentle heating at 80°C for at least 20 minutes, C-7 pteridines remained in the skin residue. These results are not explained on the basis of pteridine solubilities alone, since isoxanthopterin, which was always extracted in large quantities from ethanol-treated skins, is considerably less solu ble than either xanthopterin or 7-methyl-xanthopterin. It is well known that prolonged exposure to alkali can and does break down melano- somes and eventually melanin. Ethanol has little or no effect on either melanin or melanosomes, but heating may promote at least the breakdown of the melanosomal membrane. Thus, these investigations indicate that xanthopterin, 7-methyl-xanthopterin and leucopterin may not share the same intracellular location as the other unconjugated pteridines. Moreover, it may be that these three pteridines which share such a close metabolic relationship with pterorhodin might in fact reside within or closely associated with the same intracellular organelle as the red pigment (i.e., the melanosome).
For many years popular opinion did not favor any association of pteridines with melanophores or any cell other than xanthophores except in some unusual cases (Bagnara, 1976). However, more recent studies now report data to the contrary (Obika, 1976).
The discovery of the unusual leaf frog melanosome containing both a pteridine and melanin raises some interesting questions concern ing what kinds of selective mechanisms might exist to account for the 92
variety of pteridines synthesized and their final cellular or organellar
deposition. It also raises some interesting questions concerning the
origin and genetic potential of pigment cells in general. Certainly no
simple mechanism can be proposed to explain all of the findings both
from the literature and from this study. However, it is highly likely
that the Pachymedusa pigmentary system presents a unique opportunity
for studying just such problems.
Pteridine turnover can and should be examined in leaf frogs.
Certain questions should be asked during such studies like: Are those
pteridines synthesized during the very early development of Pachymedusa
stable, or are they broken down and replaced with time? In other words,
after a very early peak in pteridine synthesis, does the synthetic
mechanism shut down except for whatever might be needed to account for
increases in and maintenance of the overall growth of the animal? Are
new melanophores synthesized at metamorphosis, and if so, are these the
pigment cells used for pterorhodin deposition? Are old pigment cells
also used in this capacity or not at all?
To date, questions concerning the origin, role and fate of leaf
frog melanin-producing cells have not been approached. Data from these
studies demonstrate few fluctuations in either the kinds or quantities of pteridines produced throughout most of the larval development.
Significant changes occur only during and after metamorphosis. Thus, if
pteridines are only being synthesized as necessary for the maintenance
of the developing pigmentary pattern, and if the enzymes responsible for
producing pteridines were limiting their synthesis, then little or no 93 enzyme activity might be expected to be found in the skin at any given developmental stage. This could explain the inability in these studies to detect considerable or consistent levels of XDH in skins during
Pachymedusa development. Moreover, the data demonstrate that skin XDH levels do become evident, although low, during metamorphosis, and enzyme is always detectable in stage XXV, stage XXV+ and adult skins.
Xanthine Dehydrogenase and Leaf Frog Pigmentation
Exactly what evidence is there to support or refute a role for xanthine dehydrogenase in leaf frog pigmentation? From the literature it was noted that only one enzyme was even vaguely implicated in the synthetic pathway to pterorhodin, and this enzyme (XDH), is believed to occur several steps prior to the synthesis of pterorhodin itself (see
Fig. 2). Other possible enzymatic steps closer to the actual synthesis of pterorhodin have already been discussed, and in this study only the presence and activity of XDH has been confirmed.
It has always been understood that the skin itself contains all the factors necessary for complete pteridine expression. This was demonstrated in 1960 by Hama and Obika, using neural crest hanging drop cultures from urodeles. Neural crest cells, isolated before visible pigment cell differentiation had occurred, were found to differentiate both morphologically and biochemically during the culture period of 8-14 days.
This certainly was of no help in attempting to explain the situation in Pachymedusa. However, while this may be true for pre programmed chromatoblasts originating from the neural crest, what occurs after the migration, differentiation and subsequent multiplication of
these cells may no longer be entirely under the control of the cell
itself. It is conceivable on the basis of the unusual pigmentary phen
omenon occurring in leaf frogs, and its abrupt developmental appear
ance, that factors extrinsic to the pigment cells themselves may be
supplying the cues regulating the developmental alterations in gene ex
pression necessary for additional Pachymedusa pigment cell differen
tiation.
Several lines of evidence from the literature support these
suggestions, the most persuasive coming from the rosy mutant of
Drosophila. Recently, evidence that the enzyme, XDH, is synthesized in an organ of primary gene expression (i.e., some organ other than the
eye) and transported to the eyes of pupae and pharate adults has come to light. Such evidence is supported by the fact that the r£ mutant is non-autonomous such that transplanted eyes, whether mutant or wild-type in origin, always develop in correspondence with the host tissue.
Precedence for enzymes being transported through the haemolymph of insects from an organ of primary synthesis to a target organ is dis cussed in some detail by Barrett and Davidson (1975). While this precedent seems clearcut for invertebrates, there are few if any exam ples of a similar occurrence in vertebrates.
Several pieces of information from this study circumstantially support the possibility of a similar mechanism existing in leaf frogs.
First, XDH inhibition studies employing allopurinol demonstrate a role for XDH in the synthesis of pterorhodin. As pointed out previously, 95 this role is most likely an indirect one with the enzyme functioning at some step (or steps) preceding the actual synthesis of red pigment. The initial developmental enzyme data was somewhat disappointing, since little or no activity was detectable from partially purified skin ex tracts. The enzyme was shown to be clearly capable of synthesizing pteridines as well as uric acid. The quantities of enzyme present in the livers and kidneys of Pachymedusa seem excessive in tadpoles which are not known to excrete uric acid, and in adults which excrete only 5% uric acid. Moreover, the developmental pattern of the enzyme, particu larly in the liver, seems to fluctuate following a trend very similar to that reported for serum thyroxine levels in developing bullfrog larvae.
Just (1968) found that at any point during the development of bullfrog larvae, thyroxine levels in the serum vary considerably; so much so that no statistically significant decline or rise in serum hormone levels was ever recorded. However, a distinct peak in detectable serum hormone was consistently noted just prior to the onset of metamorphosis, followed by a decline towards metamorphic climax. Liver XDH in leaf frogs demonstrated a remarkably similar trend. Kidney enzyme also re vealed a similar trend but with considerably less activity in younger animals. Skin unfortunately revealed no trend and often little or no activity at all. So, if XDH is in fact responding to fluctuating hor mone levels, then the wide standard deviations reported for much of the developmental enzyme data are more easily understood.
In later attempts to better determine actual enzyme activity in skin, crude extracts were employed. Then, XDH activity became 96 consistently detectable in skin extracts, although still in very low quantities. Furthermore, as an added bonus, larval serum was assayed for XDH and found to consistently contain active enzyme. Thus, the overriding question now is to determine what the source of serum enzyme might be. Does this enzyme originate in the liver (or kidney), and can it possibly be transported to the skin and used (perhaps rapidly) for pteridine conversions? Coincidentally, at stage XXV and XXV+, serum and skin XDH levels are nearly identical (see Table VI). Also, at stage XXIII there is a dramatic rise in serum enzyme levels followed by a decline in serum XDH and subsequent gradual rise in skin XDH. While the data for skin and serum XDH are incomplete at present, these pre liminary results, particularly the undeniable presence of enzyme in serum, are certainly suggestive.
Finally, it is clear that examination of enzyme levels and skin pteridines in animals at stages younger than swimming larvae
(stage I) must be attempted. Radioactive incorporation studies during
Pachymedusa development are necessary in order to attempt to understand pteridine turnover. In addition, incorporation studies wotild add in sight into several other aspects fundamental to vertebrate pigment pattern formation, such as where pteridines are stored intracellularly, and what kinds of pteridines are stored at any particular location.
In preliminary studies utilizing leaf frog skin, melanosomes have been successfully isolated on sucrose density gradients. Thus, the question of whether melanosomes contain pigment other than melanin (and ptero- rhodin in this case) can at last be approached. The apparent advantage 97 here ia that leaf frog skin might provide a convenient source of la belled pteridines. This, in turn, could provide the extremely sensitive assay necessary for examining the small quantities of melanosomes iso lated from developing skin.
Unfortunately, the discussion of these results appears to raise more questions concerning leaf frog pigmentation than it offers answers.
The complexity of the various aspects of this study as already discussed warrant summarizing as follows:
(1) Thyroxine may or may not play an important role in the synthesis and deposition of pterorhodin. It seems likely that if thy roxine is necessary for metamorphic pigmentary events in Pachymedusa, then the direct effect is one, as yet only partially explored but prob ably involving the synthesis of 7-methyl-xanthopterin and/or the possi ble transport of XDH to skin during metamorphosis.
The developmental responses of XDH, particularly in the liver and kidney, are candidates for regulation by thyroxine. However, this regulation may be one similar to that reported for the induction of urea cycle enzymes in other amphibians during metamorphosis (Frieden and
Just, 1970). Since leaf frogs also excrete some uric acid as adults and since XDH is the key enzyme responsible for uric acid production, then a metamorphic response to thyroxine on this basis alone may account for the rise and fall in tissue XDH levels.
(2) Three pteridines, 7-methyl-xanthopterin, leucopterin and pterorhodin, are reported for the first time to occur in a vertebrate. 98
Chrysopterin and to some extent, pterorhodin, are generally believed to be artifacts produced as a result of pteridine extraction procedures.
In this case, leucopterin is more likely the degradative artifact, and chrysopterin and pterorhodin are likely to be the naturally occurring pigments.
(3) Developmentally, 7-methyl-xanthopterin appears at a signif icant point during leaf frog metamorphosis. This pteridine, while not a limiting factor in pterorhodin synthesis, is probably a precursor of red pigment along with xanthopterin which is present throughout develop ment but increases quantitatively during metamorphosis. The abrupt appearance of pterorhodin at stage XXV may be limited by either an enzyme(s) responsible directly for pterorhodin synthesis appearing for the first time, or perhaps by the sudden induction of a transport mech anism making precursors available to the site of pterorhodin synthesis for the first time.
(4) The results of portions of this study support the views of
Parisi et al. (1976), that biopterin precedes sepiapterin in the meta bolic pathway, and that both of these precede the drosopterins. While biopterin is unquestionably present in extracts of Pachymedusa dacni- color skin, sepiapterin has not been found under any circumstances, nor are there any traces of the drosopterins. While indirect, this infor mation certainly suggests that biopterin synthesis precedes that of sepiapterin. It seems now that the next logical step should be the search for the enzyme(s) responsible for the synthesis of biopterin.
An enzyme system responsible for the conversion of dihydroneopterin 99
triphosphate to biopterin has in fact been isolated from Syrian golden
hamster kidneys (Eto et al., 1976; see Fig. 2). Unfortunately, this
group was unable to demonstrate whether or not sepiapterin was an in
termediate in biopterin synthesis since the enzyme preparation con
tained considerable sepiapterin reductase contamination. They did
however, demonstrate that the side chain of biopterin was derived from
dihydroneopterin triphosphate and not from some other compound incor
porated later along the metabolic pathway. Pachymedusa, or a similar
sepiapterin-lacking/biopterin-containing organism would be the system
of choice for determining the enzymatic mechanism responsible for the
synthesis of biopterin.
(5) Results from these studies also fully support the belief that pterorhodin resides at the end of the C-7 pteridine metabolic chain (see Fig. 2). That pterorhodin might be synthesized from ery- thropterin (Pfleiderer, 1964) is certainly not a possibility in Pachy medusa, since never, under any circumstances, was erythropterin ever detected. More likely, pterorhodin is synthesized from some combination of xanthopterin and 7-methyl-xanthopterin or possibly leucopterin.
(6) Inhibition studies of XDH suggest that pteridines are very stable and turnover is very slow and/or that the competitive inhibition of XDH by allopurinol is "leaky", allowing some XDH-sensitive pteridines to continue to be synthesized at a reduced rate. Nevertheless, such studies demonstrate, for the first time, that pterorhodin is reliant upon XDH expression for its subsequent synthesis. .00
(7) Leaf frog XDH is not unlike other XDH's from the standpoint of intracellular location, relatively high molecular weight and migra tion as a single homogeneous protein band in SDS-polyacrylamide gels, perhaps consisting of several electrophoretically identical subunits.
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