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Deoxyribonuclease activities in the development of the leopard frog, Rana pipiens
DEVELOPMENTAL
UIOLOGY,
5, 232-251
Deoxyribonuclease of the
Activities Leopard JOHN
Department
of
( 1962)
Embryology, Baltimore Accepted
Frog, R.
in the Rana
Development pipiensl
COLEMAN~,”
Carnegie lnstitution 10, Maryland May
of
Washington,
1, 1962
INTKODUCTION
Enzymes are becoming increasingly useful as indexes of embryonic differentiation, but the extent to which they play causal roles in differentiative processes remains obscure. This obscurity probably reflects the complexity of the chemical and physical events leading to cellular changes that can be regarded as differentiations. Elucidation of the factors that regulate levels of enzyme activity or rates of synthesis must be preceded by determinations of the normal developmental changes undergone by the enzymes being studied. A number of investigators have engaged in such descriptive studies. The activities of cytochrome c reductases, cytochrome oxidase, and enzymes requiring pyridine nucleotides as cofactors have recently been studied in the developing leopard frog, Rana pipiens, by Lang and Grant (1961) and Wallace (1961). Developmental studies of enzyme activities in other amphibians have been reviewed by L@vtrup (1959) and Urbani (1957). In the work reported below, the development of deoxyribonuclease activity in the Rana pipiens embryo is described. Deoxyribonucleases, as studied in mammals, generally fall into two ‘This work was carried out degree of Doctor of Philosophy ’ Predoctoral Fellow of the Health Service. 3 Present address: Institute Storrs, Connecticut.
in partial fulfill merit of the requirements for the from The Johns Hopkins University. National Cancer Institute, United States Public of
Cellular 232
Biology,
University
of
Connecticut,
DNASE
ACTIVITIES
IS
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DEVELOPMEN
233
categories designated deoxyribonuclease (DNase) I1 and DNase II (cf. Laskowski, 1961). These two categories are distinguished by their pH optima, ionic requirements, and the specificity of the phosphodiester bonds which they cleave, DNase I forming 5’-nucleotides and DNase II forming 3’-nucleotides. It must be emphasized that the terms DNase I and DNase II as used herein refer only to the enzymatic breakdown of DNA under the conditions set forth and do not necessarily imply identity with the purified DNase I and DNase II studied in mammals. Deoxyribonuclease activities have previously been studied in Rana pipiens embryos by Finamore (1955) and Blumenthal (1957). Finamore reported low cyclical variations in the activity of a DNase I in the embryo up to the early neurula. Employing a different assay technique, Blumenthal was unable to confirm Finamore’s findings. She found no DNase I activity in embryonic, larval, or adult frogs, and could detect DNase II activity only in late tadpoles and adults. The present investigation was undertaken in an attempt to resolve these differences by a systematic determination of optimal conditions of assay. Both types of DNase activity have been studied in tadpoles and adult frogs, the developmental stages at which each type of activity increases have been defined, and the possible functional significance of this timing has been examined. A preliminary account of this work has been published (Coleman, 1961). MATERIALS
Biological
AND
METHODS
Material
Adult Rana pipiens were stored in a cold room maintained at 4°C. Induction of ovulation, stripping, and insemination were carried out essentially as described by Hamburger (1942) except that pituitaries were macerated prior to injection. Developing embryos and tadpoles were kept at 21°C in 10% Holtfreter’s solution supplemented with lo-’ hl MgSO,. Tadpoles were fed a commercially prepared diet of dried vegetable matter, “Nature Flakes.” ‘Abbreviations used in this paper: DNA, deoxyribonucleic acid; DNase I, alkaline deoxyribonuclease; DNase II, acid deoxyribonuclease; RNase, ribonuclease; EDTA, di-sodium ethylenediamine tetraacetate; Tris, 2-amino-2-hydroxymethyl-l,%propanediol; PCA, perchloric acid; BSA, bovine serum albumin.
234
JOHN
Preparation
1% COLEMAN
of Homogenates
Homogenates were prepared by grinding samples in glass Ten Broeck homogenizers in chilled distilled water containing 1 mg bovine serum albumin (BSA) per milliliter. After standing in an ice bath for approximately 30 minutes with occasional rehomogenization, they were centrifuged for 10 minutes at 2500 g in an International refrigerated centrifuge, model PR-2, to remove debris. Homogenates were kept in an ice bath until used. Dilutions of the homogenate were prepared using the BSA solution as a diluent, the BSA being present to protect the enzyme activity at higher dilutions. The jelly layers of unfertilized eggs and prehatching embryos were removed manually with fine jewelers’ forceps before homogenization. Posthatching larvae and adult tissues were homogenized directly. Wet weights of tadpoles and adult tissues were recorded, and samples of the homogenates were retained for protein determinations as described below. For early developmental stages, ten embryos were homogenized per milliliter of the BSA solution. In later stages the concentration was dictated by the amount of enzyme activity present. Protein
Determinations
Protein was determined by the method of Lowry et al. (1951) modified to permit assay of a 10-J sample of homogenate. A standard curve consisting of several concentrations of BSA was run with each set of determinations. Enzyme
Assays
The assay used for both DNase I and DNase II depends upon the insolubility of macromolecular DNA in cold 2% perchloric acid (PCA). Ten milliliters of appropriate substrate solution was placed in 25-ml Erlenmeyer flasks fitted with ground-glass stoppers, clamped to a Burrell wrist-action shaker, and oscillated gently in a constanttemperature water bath at 27°C. To start the reaction 2 ml of homogenate was added; at 0, 30, 60, and 120 minutes, l-ml aliquots were pipetted in duplicate into chilled tubes, each containing 0.2 ml of cold 12% PCA, mixed thoroughly, and allowed to stand in an ice bath for 15 minutes. The precipitate was sedimented by centrifugation at 2000 g for 10 minutes in an International refrigerated centrifuge. The optical density of the supernatant fluid, containing oligonucleotides
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235
resulting from the degradation of DNA, was read at 260 rnp on the Beckman DU spectrophotometer against an appropriate blank. Ultraviolet absorption spectra of the PCA supernatants were determined on the Beckman DK-2 recording spectrophotometer. One unit of DNase activity is defined as that amount of activity which causes an increase in absorbancy at 260 rnp of 0.001 per 30 minutes of incubation at 27°C. The pH of each flask was determined at the end of each experiment. Substrate solutions were made up fresh every 2 or 3 days, and the DNA stock solution (2 mg DNA per milliliter in distilled water) approximately every week. Validity of the method was initially ascertained by using partially purified commercial DNase (Worthington Biochemical Corporation). Highly purified salmon sperm DNA (California Corporation for Biochemical Research) was used as substrate throughout. Buffers were acetic acid-sodium acetate, Tris-hydrochloric acid, or glycine-sodium hydroxide, as specified in the text. To study the development of DNase activities, approximately equal numbers of eggs were taken from each of four gravid females, fertilized with the same sperm suspension, and sampled at daily intervals until Shumway stage 25 was attained. Determinations were made at approximately the same time every day. Reaction flasks were set up for each enzyme determination as follows: 1. 2. 3. 4.
Homogenate plus buffer BSA plus substrate Homogenate plus substrate Homogenate plus substrate plus inhibitor DNase I inhibited by 0.01 M EDTA DNase II inhibited by 0.05 M MgSO,
Because this investigation involved the use of crude homogenates of developing embryos in which unknown phosphodiesterase activities might be present, it was considered desirable to include a more direct control than that provided by summation of flasks nos. 1 and 2. Thus flask no. 4 was designed to measure the maximal amount of hydrolytic activity that could occur when the activity of DNase I or II was inhibited. When it was expedient to do so, flask no. 4 was set up as a duplicate of flask no. 3. This invariably led to very close duplication of results.
236
JOHN
R. COLEMAN
RESULTS
Before investigating the development of DNase activities in ontogenesis, it was necessary to attempt to define conditions that would be optimal for enzyme assay. DNase
I
Homogenates of tadpoles beyond frog pancreas were found to contain
Shumway stage 25 and adult a DNase activity that required
4000 A
Frog
pancreas
0
Frog
liver
DNase
DNase
6.0
7.0
I
1I
z w
3000
k cc a F L a? u)
2000
s 0 ?
IO00
z 3
1
5.0
0.0
9.0
10.0
PH FIG. 1. Effect of pH on DNase activities of adult frog tissue homogenates. Composition of substrate for DNase I: pH 5.0-6.0, 0.025 M acetate buffer; pH 7.0-9.0, 0.025 M Tris-HCl buffer; pH 9.5, 0.025 M glycine-NaOH buffer; all contained 08.025 M MgS04 and 0.2 mg DNA per milliliter. Composition of substrate for DNase II: pH 4.0-6.3, 0.1 M acetate buffer; pH 7.0-9.0, 0.1 M TrisHCl buffer; all contained 0.1 M NaCI, 0.001 M EDTA, and 0.2 mg DNA per milliliter. Concentration of homogenate for DNase I was 0.07 mg protein per milliliter, for DNase II was 0.14 mg protein per milliliter.
DNASE
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DEVELOPMENT
400 DNose o DNose
A
I n
300 -i \ W
cn
I
0
0.1
0.2
mg PROTEIN
0.3
/ml
0.4
0.5
0.6
0.7
HOMOGENATE
FIG. 2. Effect of homogenate dilution on tadpole DNase activities. Composition of substrate for DNase I: 0.025 M Tris-HCl, pH 8.6, 0.005 M MgSO,, 0.2 mg DNA per milliliter; for DNase II: 0.3 M acetate buffer, pH 4.8, 0.001 M EDTA, 0.2 mg DNA per milliliter. DNase I curve is from a whole 36-mm tadpole, Taylor and Kollros stage 4; DNase II curve is from a Taylor and Kollros stage 13 tadpole tail.
magnesium and exhibited optimal activity at pH 8.59.0 in 0.025 M Tris-HCI buffer (Fig. 1) . This activity was designated DNase I. At a DNA concentration of 0.2 mg per milliliter, the optimal MgSO, concentration was found to be in the range of 0.0054025 M, varying slightly with the preparation. Concentrations of MgSO, above 0.025 M were increasingly inhibitory. Omitting MgSO, from the substrate medium reduced the activity to about 20% of that found in the presence of 0.005 M MgSO,. That this requirement is for divalent cations was indicated by the finding that a chelating agent, ethylenediamine tetraacetate (EDTA ) , was capable of inhibiting the reaction. As
238
JOHN
R. COLEMAN
little as 0.0005 M EDTA reduced the activity to about 2% of that found in the presence of magnesium. This inhibition could be overcome by increasing the concentration of MgSO,. When the specific activity of tadpole DNase I was plotted as a function of the DNA concentration in 0.025 M MgSO*, saturation with substrate was found at a concentration of 0.2 mg DNA per milliliter. Therefore, at this substrate concentration, the amount of activity observed during the 2-hour incubation should be directly proportional to the concentration of enzyme. Determinations of the DNase I activity in various dilutions of the homogenate showed that this was the case (Fig. 2).
WAVELENGTH
mp
FIG. 3. Absorption spectra of PCA supernatant digestion of DNA by tadpole DNase I. Composition HCl buffer, pH 8.7, 0.005 M MgSO,, 0.2 mg DNA of homogenate was 0.16 mg protein per milliliter. stage 25.
solutions during course of of substrate: 0.025 M Trisper milliliter. Concentration Tadpoles were Shumway
controls in Ultraviolet absorption spectra, as well as appropriate which DNA or homogenate was omitted from the reaction mixture, indicated that the reaction observed with this assay technique was breakdown of DNA following its exposure to an appropriate homogenate (Fig. 3). Table 1 shows the substrate composition which was found to be optimal for the assay of frog DNase I activity.
DNASE
ACTIVITIES
IN
TABLE
FROG
239
DEVELOPMENT
1
COMPOSITION OF SUBSTRATES FOR DEVELOPMENTAL STUXES DNase I
DNase II
0. u25 &I Tris-HCl 0.005 XI MgS04 0.2 mg DNA per ml pH 8.7 2~ 0.1 p = 0.03
U. 3 AI acetate 0.001 M EDTh 0.2 mg DNA per ml pH 4.9 x!z 0.1 p = 0.18
DNase II Another DNase, designated DNase II, was found in adult tissue and tadpole homogenates. This activity exhibited an optimum at pH 4.5-5.0 at all stages tested (Shumway stage 19 embryo, 3-month-old tadpole, adult frog liver) (Fig. 1). DNase II does not require divalent ions, but is inhibited by them. Thus, a concentration of MgSO, of 0.01 M inhibited the activity 95%. On the other hand, optimal activity was exhibited at an ionic strength ( ,u) of 0.18 in the presence of monovalent ions as provided by acetic acid-Na acetate buffer, or by mixtures of acetate buffer and NaCl. As recommended by Laskowski ( 1961), a small amount of EDTA (0.001 M ) was included in the reaction mixture to prevent inhibition of DNase II activity by divalent cations in the crude homogenates (Table 1). As the concentration of DNA in the reaction mixture was increased, it was found that activity decreased, i.e., substrate inhibition was encountered. At a DNA concentration of 0.2 mg/ml, the homogenate dilution curve was linear (Fig. 2), indicating that the amount of activity observed was proportional to the concentration of enzyme. Substrate inhibition was apparent only in the failure of the curve to bisect the origin. Since substrate became limiting too rapidly at lower DNA concentrations, 0.2 mg DNA per milliliter was selected for use in the developmental studies (Table 1). As in the case of DNase I activity, ultraviolet absorption spectra and appropriate controls indicated that the reaction being followed was the hydrolysis of DNA. For both DNase I and DNase II activities, the increase in absorbancy of the PCA supernatant solution was not a strictly linear function
240
JOHN
R. COLEMAN
of the time of incubation (Fig. 4). The rate of appearance of acidsoluble material absorbing at 260 rnp increased with the time of incubation until the substrate concentration became limiting, after which the rate decreased, resulting in a sigmoid curve. In spite of this, the linearity of the homogenate dilution curves derived from the 60-120 minute interval of incubation (Fig. 2) justified a quanti,600 n
DNose
I
0
DNose
H
: N ti
m u
,400
0
30 TIME
60 IN
90
I20
MINUTES
FIG. 4. Progression of DNA digestion by larval homogenates. of substrates same as for Fig. 2. Concentration of homogenate for 1.1 mg protein per milliliter; for DN ase II, 0.44 mg protein Larvae were early Shumway stage 25.
Composition DNase I was per milliliter.
tative consideration of the amount of nuclease activity present. For technical reasons, the 0 time values were occasionally erratic and initial reaction rates could not be accurately estimated. Consequently, the units of DNase activity reported herein refer to the portion of the curve obtained between 60 and 120 minutes of incubation. Comparable determinations with commercial bovine pancreatic DNase I gave similar results. Dezjelopment
of DNase Activities
During the determination of optimal conditions for enzyme assay, it was found that both DNase activities were low in early embryonic stages, but present in tadpoles and adults. On this basis, it was decided to undertake a rigorous examination of both types of activity
DNASE
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241
simultaneously in developing embryos until both activities were present in significant amounts. Under the conditions employed, DNase I activity was essentially undetectable prior to stage 23, the beginning of opercular fold formation (Figs. 5 and 6). Activity began to increase slightly at stage 23, 800
700
5
600-
& LL Q.
500-
-
: \
400
-
300
-
200
-
: z cl if z 3
FG
N 18 20
STAGE
22
OF
24
25
Todpolep
DEVELOPMENT
FIG. 5. DNase I activity per milligram protein of the frog. Substrate composition was as shown Development is expressed in Shumway stages.
during early development in Table 1 for DNase
I.
but did not increase rapidly until the operculum was closing at stage 24 or early stage 25. On a protein basis (Fig. 5), the activity appeared to reach a peak and then decline, but the data were quite variable in this region of the curve. On a per embryo basis, this same peak was observed, but activity thereafter continued to increase rather than level off or drop (Fig. 6). From its first appearance at stage 23, the nuclease activity was 90-95% inhibitable by 0.01 M EDTA. A very low level of DNase II activity was found in the unfertilized
242
JOHN
R. COLEMAN
egg, and this dropped slightly through gastrulation (Figs. 7 and 8). With the advent of neurulation, however, this activity began to increase and, on a per milligram protein basis, continued to increase until about stage 24, when it began to level off and perhaps declined
0
F G N 18
STAGE
FIG. 6. Same data
20
22
OF
DNase I activity per embryo as those plotted in Fig. 5, but
24
25
Tadpoles-
DEVELOPMENT
during early development of the frog. calculated on a per embryo basis.
somewhat (Fig. 7). On a per embryo basis, such a change in rate was observed only as a slight shoulder in the curve, after which the activity continued to increase (Fig. 8). The activity observed in these determinations was completely inhibitable by 0.05 M MgSO,. Since the rapid increase in DNase I activity at early stage 25 was approximately coincident with the onset of feeding, it was considered that this activity might be localized in the digestive tract. Stage 25 tadpoles were grown for 1 week to a length of 17 mm, dissected in cold isotonic saline, and the component parts assayed for DNase I activity (Table 2). Gut had some activity, but pancreas was found to be by far the most active tissue examined. DNase II was not com-
F G N
18
20 22
STAGE
24
OF
25
Tadpole-
DEVELOPMENT
FIG. 7. DNase II activity per milligram protein of the frog. Substrate composition was as shown Development is expressed in Shumway stages.
in
during Table
early development 1 for DNase II.
600
F Lx !ii w \
500
400
2 D fi
300
c z 1
200
100
F G N I8
STAGE
20
22
OF
24
25
Tadpoles-
DEVELOPMENT
FIG. 8. DNase II activity per embryo during early development frog. Same data as those plotted in Fig. 7, but calculated on a per embryo Development is expressed in Shumway stages. 243
of
the basis.
244
JOHN
parably localized, although than in the body (Table pancreas.
H. COLEYIAN
its specific activity was higher in the gut 2). In this case, gut included liver and TABLE 2
DISTRIBUTIOK ACTIVITIES
IN
Units Preparation
OF DNASE
IlNnse
Embryo
Whole Gut
SHUMWAY
Pancreas
DNASE
AXD
25
II
TADPOLIW
I *er:
l:nits
DNrtse
II per:
Mg protein
Embryo
406 103 1830 20 !I
689 270
:w1 500
32
350
288 68 330 2 6
Liver Body
I
STAGE
Mg protrin
a Tadpoles were grown to a length of 17 mm and dissected in cold isotonic saline. The digestive tract, with liver and pancreas attached, was separated from the rest of the body after severing the gut immediately anterior t,o the stomach and at the opening into the cloaca. The portion remaining behind is designated “body.” For DNase I, “gut” represents the digestive tract from which liver and pancreas have been removed and assayed separately. For DNase II, “grit” includes liver and pancreas. “Whole” represents undissected tadpoles.
Activities were too low and the embryonic systems were changing too rapidly to permit comparable dissections designed to localize the early increase in DNase II activity. It has been hypothesized that TABLE DNASE
II AND
(Taylor
IICTIVITY
3
TAILS
NONMETAMORPHOSING
stage and liollrou)
13
13 18 2lY 22 n Beginning
IN
Units DNa8c II per mg protein
348 395 705
OF METAMORPHOSIW TAI)POLES l!nitti DKase II per rng wet weight
8.0 8.6 18.3
1170
31.7
1160
il.8
of tail resorption.
this hydrolytic enzyme may play a role in degenerative processes (cf. de Duve, 1959; Novikoff, 1961). To investigate its correlation with a degenerative process in the frog, the DNase II activities of homogenates of tadpole tails were determined before and during resorption
DNASE
ACTIVITlES
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945
of the tail at metamorphosis. Tails in the process of resorption had severalfold the DNase II activity of tails not undergoing resorption (Table 3). DISCUSSION
The failure to detect DNase I activity in the early frog embryo reported in this work is in apparent conflict with the results of Finamore (1955). However, the activities reported by Finamore were very low, were obtained at suboptimal pH (pH 7.4), and apparently were not controlled for homogenate autolysis. Hence, his results do not necessarily reflect nuclease activity. Blumenthal (1957) was unable to detect DNase I activity in embryos, tadpoles, or adults of RUMZ pipiens. In this case, failure to detect activity was probably due to the use of suboptimal pH (pH 4-7) in combination with high ionic strength (0.2 M phosphate buffer) leading to complete inactivation of DNase I (cf. Rotherham et al., 1956). Blumenthal found DNase II activity in adult frogs, but was unable to detect it in embryos or tadpoles. These results led her to suggest that this enzyme probably appeared at metamorphosis. In the present investigation, DNase II activity was detected at all stages of development, although activity was very low prior to neurulation. In Blumenthal’s work, it is likely that a high substrate concentration (1.5 mg DNA per milliliter) in combination with low ionic strength (0.1 M acetate buffer, ,U approximately 0.06) prevented observation of DNase II activity in any but the most active preparations. The determination of optimal assay conditions permits, as a corollary, comparison of the reaction characteristics of frog DNase I and DNase II activities with the more extensive information available on mammalian DNases (reviewed by Laskowski, 1961). Except for the pH optimum of frog DNase I activity, other characteristics of the DNase activities reported here are quite similar to those of the DNase I and DNase II activities studied in mammals. In fact, the conditions for the assay of frog DNase II activity derived from the investigations reported here are very close to those recommended by Laskowski (1961) for the study of DNase II in higher vertebrate tissues. The exceptionally alkaline pH optimum of frog DNase I activity may represent a real divergence from the type of DNase I activity found in higher forms or may be a function of the use of crude homogenates. Siebert et al. (1950) reported considerable variation in the pH opti-
mum for DNase activity in crude homogenates depending on the tissue and animal investigated. Substrate inhibition is commonly encountered in studies of both DNase I and DNase II (0th et al., 1958; Laskowski, 1961) although, in the present investigation, it was encountered only with DNase II activity. The extent to which this may reflect the presence of divalent cations in the DNA preparation is not clear, but 0th et al. report no change in activity after dialysis of DNA against water or EDTA. The linearity of the homogenate dilution curves (Fig. 2) justifies the consideration of DNase activities in quantitative terms in spite of substrate inhibition and the nonlinear rate of appearance of reaction products (Fig. 4). The sigmoid relationship of the appearance of acid-soluble products with time of incubation is intimately related to the endonuclease nature of these enzymes, rather than being a consequence of the use of suboptimal conditions and crude homogenates. Similarly complex characteristics seem to prevail even in studies of purified DNase I (Kunitz, 1950) and DNase II (see Laskowski, 1961, for detailed discussion). Comparison of DNase I and DNase II activities in the frog embryo shows that they undergo strikingly different patterns of development (Figs. 5-B). The initial rise in DNase II activity approximately coincides with neurulation, occurring much earlier and at a slower rate than the DNase I activity. DNase I begins to increase during at which time the yolk reserves are beopercular fold formation, coming depleted, and approaches maximal activity at about the time active feeding begins at Shumway stage 25. On a protein basis, the increase in DNase II activity ceases at about the time DNase I activity becomes detectable (Figs. 5 and 7). The development of DNase I activity between stages 23 and 25 can be correlated rather closely with the morphological development of the amphibian pancreas as reported by other investigators (Huettner, 1949; Rugh, 1951; Tahara and Nakamura, 1961) and, in fact, this activity may be localized in the pancreas (Table 2). In investigations on mammals, the detection of DNase I activity in tissues other than pancreas is frequently complicated by inhibitors (Dabowska et al., 1949; Cunningham and Laskowski, 1953; Feinstein and Hagen, 1959). The relatively high activity of the isolated pancreas compared with the activity of the whole tadpole (Table 2) may reflect
DNASE
ACTIVITIES
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DEVELOPMENT
247
the presence of such inhibitors in tadpole tissues, although no critical attempts to demonstrate them have been made. The correlation between the development of DNase I activity and the onset of active feeding at stage 25 lends support to the hypothesis of Allfrey and Mirsky (1952) that the function of pancreatic DNase I is primarily digestive. The DNase I activity found in gut homogenates (Table 2) may thus be of pancreatic origin. Relatively few of the enzymes studied in anuran embryos increase as late in embryonic development as does DNase I activity. Those which do include a-amylase (Scollo-Lavizzari, 1956), an alkaline protease, and lipase in Bufo culgaris (Urbani, 1957), an alkaline protease in Xenopus (Lovtrup, 1955), and DPNH-cytochrome c reductase in Rana pipiens (Lang and Grant, 1961; Wallace, 1961). No attempts at tissue localization have been reported for these enzymes, but at least the first four mentioned could reflect pancreatic development. In contrast with DNase I, DNase II activity began to increase during neurulation; at this time localized degenerative processes have been reported to occur in the formation of the neural plate and neural tube (Glucksmann, 1951). In mammals, DNase II has been shown to be compartmentalized with other acid hydrolases in the lysosomal fraction of tissue homogenates (de Duve et al., 1955; van Lancker and Holtzer, 1959) and has been hypothesized to play a degradative role in regions of cellular degeneration (see de Duve, 1959, and Novikoff, 1961, for extensive reviews). The increase in DNase II activity in the early neurula may reflect such degenerative processes, but for technical reasons no attempt was made to determine the regional distribution of DNase II activity in the early embryo. By Shumway stage 25, the activity appears to be widespread (Table 2). Of other acid hydrolases studied in anuran embryos, acid protease ( Urbani, 1957)) acid phosphatase and acid RNase (de Cesaris Coromaldi, 1955, 1958) begin to increase in the tailbud stage of Bufo uulgaris, whereas cathepsin increases in the gastrula stage of Xenopus Eaewis (Deuchar, 1958). Information concerning the extent to which these developmental differences might reflect differences in compartmentalization is currently lacking. An .attempt was made to correlate the activity of DNase II with a more accessible degenerative process, the resorption of the tadpole
248
JOHN
H. COLEMAN
tail at metamorphosis. On a protein basis, tails in the process of resorption were found to have about three times the DNase II activity of nonresorbing tails (Table 3). On a wet-weight basis, this figure is enlarged to fivefold. These results are in close agreement with those of other workers on hydrolytic enzymes in anuran metamorphosis (Urbani, 1957; Weber, 1957; de Cesaris Coromaldi, 1958; Weber and Niehus, 1961) and have been confirmed in more extensive investigations of thyroxine-induced tail resorption in R. catesbeiana (Coleman, unpublished). Finally, no critical information is available concerning the extent to which the increases in nuclease activities reported in this paper reflect de novo synthesis of new enzyme protein. Preliminary experiments with mixtures of homogenates from different stages of development indicate that inhibitors and/or activators may not play a major role in the regulation of the observed DNase activities. Since DNase II activity increases at a time when the embryo depends completely on its stored yolk reserves, more sophisticated biochemical approaches are difficult. On the other hand, the relatively late increase in DNase I activity may permit investigation of this question through the use of radioisotopes or immunological cross reactivity, pending preliminary purification of the enzyme proteins involved. Until the problem of de now synthesis is clarified, little can be said about the mechanisms that regulate enzyme activity or synthesis. SUMMARY
Deoxyribonucleases have been studied in the leopard frog, Rana pipiens, by following the progressive increase of perchloric acidsoluble compounds absorbing at 260 rnp during incubation of homogenates with macromolecular DNA at 27°C. The terms DNase I and DNase II, as used in this report, do not necessarily imply identity with purified DNase I and DNase II as studied in mammals, but refer only to the enzymatic breakdown of DNA under two given sets of conditions. Frog DNase I activity exhibits an alkaline pH optimum of pH 8.5-9.0, but in other requirements is similar to DNase I activities studied in higher vertebrates. It is relatively unaffected by excess substrate, exhibits a divalent cation requirement which can be satisfied by magnesium but is inhibited by concentrations of MgSO, above 0,025 M, Frog DNase II activity is similar to mammalian DNase II
DNASE
ACTIVITIES
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249
DEVELOPMENT
in all the requirements studied. It has a pH optimum of 4.5-5.0, exhibits a high degree of sensitivity to excess substrate, requires an ionic strength of about 0.18 for optimal activity, and is strongly inhibited by divalent ions. DNase I activity was found in adult pancreas and whole tadpole homogenates. By the assay technique employed, it was undetectable in the early embryo but began to rise at approximately Shumway stage 23. This coincides approximately with the time of morphological appearance of the anuran pancreas as reported in the literature. Stage 25 tadpoles grown for 1 week and dissected showed a striking localization of DNase I activity in the pancreas, and perhaps some in the gut as well. Low DNase II activity was found in the unfertilized egg. This activity decreased slightly through gastrulation, then began to increase with the onset of neurulation. On a per milligram protein basis, this activity continued to increase through early prelarval development but leveled off at about the time DNase I activity became detectable. On a per embryo basis, the activity continued to increase. Hence, these two nuclease activities show strikingly different patterns of development. The increase in DNase II activity at neurulation coincides with cellular degeneration reported to occur in the forming neural plate and neural tube of vertebrate embryos. No attempt was made to determine the localization of DNase II activity in the neurula. A three- to fivefold increase in activity was found during tail resorption at metamorphosis, a result which suggests a correlation between DNase II activity and degenerative processes. The author gratefully acknowledges the persistent encouragement tion of Dr. James D. Ebert, Dr. Yoshihiro Kato, and Dr. Annette Acknowledgment is also extended to Dr. Heinz Herrmann for his of the manuscript.
and
stimulaColeman. critical reading W.
REFERENCES
ALLFHEY, V., and MIRSKY, A. E. ( 1952).
Some aspects of the desoxyribonuclease of animal tissues. J. Gen. Physiol. 36, 227-241. BLUMENTHAL, G. ( 1957). A comparative study of the desoxyribonuclease activity in adult and embryonic tissue. J. Embryol. Exptl. Morphol. 5, 377-395. COLEMAN, J. R. ( 1961). Deoxyribonucleases in embryonic and adult Rana pipiens. (Abstract) Am. Zoologist 1, 348-349, CUNNINGHAM, L., and LASKQWSIU, M. (1953). Presence of two different desoxyactivities
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JOHN
R. COLEMAiY
ribonucleodepolymerases in veal kidney. Biochim. et Biophys. Acta 11, 590-591. DABOWSKA, W., COOPER, E. J., and LASKOWSKI, M. ( 1949). A specific inhibitor for desoxyribonuclease. J. Biol. Chem. 177, 991-992. DE CESARIS COHOMALDI, L. ( 1955). Studio della fosfatasi nello sviluppo embrionale e larvale di Bufo vulguris. Ricerca xi. 25, 2323-2333. DE CESARIS COROMALDI, L. ( 1958). Studio della ribonucleasi nello sviluppo embrionale, larvale e nella metamorfosi di Bufo uulgaris. Rend. ist. lombn/do sci. e lettere B92, 357-378. I)E DUVE, C. ( 1959). Lysosomes, a new group of cytoplasmic particles. In “Sulk cellular Particles” (T. Hayashi, ed.), pp. 128-159. Ronald, New York. DE DUVE, C., PRESSMAN, B. C., GIANETTO, R., WATTIAUX, R., and APPEL~ZANS, F. ( 1955). Tissue fractionation studies. 6. Intracellular distribution patterns of enzymes in rat liver tissue. Biochem. J. 60, 604-617. DEUCHAR, E. M. (1958). Regional differences in catheptic activity in Xenopus laevis embryos. J. Embryol. Exptl. Morphol. 6, 223-237. FEINSTEIN, R. N., and H,GEN, U. ( 1959). Tissue distribution of the neutral deoxyribonuclease ( DNase I). ( Abstract) Federation PTOC. 18, 225. FINAMORE, F. J. ( 1955). A study of ribonuclease and desoxyribonuclease during the early developmental stages of Rana pipiens. Exptl. Cell Research 8, 533542. GL~~CKSMANN, A. (1951). Cell deaths in normal vertebrate ontogeny. Biol. Revs. Cambridge Phil. Sot. 26, 59-86. HAMBURGER, V. ( 1942). “A Manual of Experimental Embryology,” pp. 30-35. University of Chicago Press, Chicago, Illinois. HUETTNEH, A. F. ( 1949). “Comparative Embryology of the Vertebrates,” p. 134. Macmillan, New York. KUNITZ, M. (1950). Crystalline desoxyribonuclease. II. Digestion of thymus nucleic acid (desoxyribonucleic acid). The kinetics of the reaction. J. Gen. Physiol. 33, 363-377. LANG, C. A., and GHANT, P. ( 1961). Respiratory enzyme changes during frog embryogenesis. PTOC. Natl. Acad. Sci. U. S. 47, 12361244. LASKOWSKI, hl. ( 1961). Deoxyribonucleases. In “The Enzymes” (P. D. Boyer, H. Lardy, and K. Myrback, eds.), Vol. V, pp. 123-148. Academic Press, New York. L@VTHUP, S. ( 1955). Chemical differentiation during amphibian embryogenesis. Compt. rend. trav. lab. C&berg S&r. chim. 29, 261-314. LGVTRUP, S. (1959). Biochemical indices of embryonic differentiation. P~oc. lntern, Congr. Biochem. 4th Congr. Vienna 1958, Vol. 6, 105-120. LOWRY, 0. H., ROSEBROUGH, N. J., FARH, A. L., and RANDALL, R. J. (1951). Protein measurement with the Folin phenol reagent. .I. Biol. Chem. 193, 265275. NOVIKOFF, A. ( 1961). Lysosomes and related particles. In “The Cell” (J. Brachet and A. E. Mirsky, eds.), Vol. 2, pp. 423-488. Academic Press, New York. OTH, A., FREDERICQ, E., and HACHA, R. (1958). Enzymic degradation of deoxyribonucleic acid. I. Purification of acid deoxyribonuclease. Biochim. et Biophys. Acta 29, 287-296. ROTHERHAM, J., SCHOTTELIT D. D., IRVIN, J, L., and IRVIN, E. M. (1956).
DNASE
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The deoxyribonuclease of rat liver in relation to the isolation of deoxyribonucleoprotein. J. Biol. Chem. 223, 817-829. RUGH, R. ( 1951). “The Frog. Its Reproduction and Development,” p. 204. Blakiston, Philadelphia. SCOLLO-LAVIZZARI, G. ( 1956). I1 comportamento delle alfa e beta amilasi durante lo sviluppo embrionale e larvale di Bufo vulguris. Ricerca sci. 26, 2146-2152. SHUMWAY, W. ( 1940). Stages in the normal development of Rana pipiens. I. External form. Anat. Record 78, 139-147. SIEBERT, G., LANG, K., and CORBET, A. ( 1950). Uber das pH-Optimum der Desoxyribonuclease verschiedener Tierarten. Biochem. 2. 320, 418421. TAHARA, Y., and NAKAMURA, 0. (1961). Topography of the presumptive rudiments in the endoderm of the anuran neurula. J. Embryol. Exptl. A40rph0l. 9, 138-158. TAYLOR, A. C., and KOLLROS, J. J. (1946). Stages in the normal development of Rana pipiens larvae. Anat. Record 94, 7-24. URBANI, E. (1957). Studi di embriologia e zoologia chimica degli anfibi. Rend. id. lombardo sci. e lettere B92, 69-179. VAN LANCKER, J. L., and HOLTZER, R. L. ( 1959). Tissue fractionation studies of mouse pancreas. Intracellular distribution of nitrogen, deoxyribonucleic acid, ribonucleic acid, amylase, acid phosphatase, deoxyribonuclease, and cytochrome oxidase. j. Biol. Chem. 234, 2359-2363. WALLACE, R. A. (1961). Enzymatic patterns in the developing frog embryo. Develop. Biol. 3, 486-515. WEBER, R. (1957). On the biological function of cathepsin in tail tissue of Xenopus larvae. Experientia 13, 153-155. WEBER, R., and NIEHUS, B. ( 1961). Zur Aktivitat der sauren Phosphatase im Schwanz der Xenopuslarven wahrend Wachstum und Metamorphose. Helv. Physiol. et Pharmacol. Acta 19, 103-117.
UIOLOGY,
5, 232-251
Deoxyribonuclease of the
Activities Leopard JOHN
Department
of
( 1962)
Embryology, Baltimore Accepted
Frog, R.
in the Rana
Development pipiensl
COLEMAN~,”
Carnegie lnstitution 10, Maryland May
of
Washington,
1, 1962
INTKODUCTION
Enzymes are becoming increasingly useful as indexes of embryonic differentiation, but the extent to which they play causal roles in differentiative processes remains obscure. This obscurity probably reflects the complexity of the chemical and physical events leading to cellular changes that can be regarded as differentiations. Elucidation of the factors that regulate levels of enzyme activity or rates of synthesis must be preceded by determinations of the normal developmental changes undergone by the enzymes being studied. A number of investigators have engaged in such descriptive studies. The activities of cytochrome c reductases, cytochrome oxidase, and enzymes requiring pyridine nucleotides as cofactors have recently been studied in the developing leopard frog, Rana pipiens, by Lang and Grant (1961) and Wallace (1961). Developmental studies of enzyme activities in other amphibians have been reviewed by L@vtrup (1959) and Urbani (1957). In the work reported below, the development of deoxyribonuclease activity in the Rana pipiens embryo is described. Deoxyribonucleases, as studied in mammals, generally fall into two ‘This work was carried out degree of Doctor of Philosophy ’ Predoctoral Fellow of the Health Service. 3 Present address: Institute Storrs, Connecticut.
in partial fulfill merit of the requirements for the from The Johns Hopkins University. National Cancer Institute, United States Public of
Cellular 232
Biology,
University
of
Connecticut,
DNASE
ACTIVITIES
IS
FROG
DEVELOPMEN
233
categories designated deoxyribonuclease (DNase) I1 and DNase II (cf. Laskowski, 1961). These two categories are distinguished by their pH optima, ionic requirements, and the specificity of the phosphodiester bonds which they cleave, DNase I forming 5’-nucleotides and DNase II forming 3’-nucleotides. It must be emphasized that the terms DNase I and DNase II as used herein refer only to the enzymatic breakdown of DNA under the conditions set forth and do not necessarily imply identity with the purified DNase I and DNase II studied in mammals. Deoxyribonuclease activities have previously been studied in Rana pipiens embryos by Finamore (1955) and Blumenthal (1957). Finamore reported low cyclical variations in the activity of a DNase I in the embryo up to the early neurula. Employing a different assay technique, Blumenthal was unable to confirm Finamore’s findings. She found no DNase I activity in embryonic, larval, or adult frogs, and could detect DNase II activity only in late tadpoles and adults. The present investigation was undertaken in an attempt to resolve these differences by a systematic determination of optimal conditions of assay. Both types of DNase activity have been studied in tadpoles and adult frogs, the developmental stages at which each type of activity increases have been defined, and the possible functional significance of this timing has been examined. A preliminary account of this work has been published (Coleman, 1961). MATERIALS
Biological
AND
METHODS
Material
Adult Rana pipiens were stored in a cold room maintained at 4°C. Induction of ovulation, stripping, and insemination were carried out essentially as described by Hamburger (1942) except that pituitaries were macerated prior to injection. Developing embryos and tadpoles were kept at 21°C in 10% Holtfreter’s solution supplemented with lo-’ hl MgSO,. Tadpoles were fed a commercially prepared diet of dried vegetable matter, “Nature Flakes.” ‘Abbreviations used in this paper: DNA, deoxyribonucleic acid; DNase I, alkaline deoxyribonuclease; DNase II, acid deoxyribonuclease; RNase, ribonuclease; EDTA, di-sodium ethylenediamine tetraacetate; Tris, 2-amino-2-hydroxymethyl-l,%propanediol; PCA, perchloric acid; BSA, bovine serum albumin.
234
JOHN
Preparation
1% COLEMAN
of Homogenates
Homogenates were prepared by grinding samples in glass Ten Broeck homogenizers in chilled distilled water containing 1 mg bovine serum albumin (BSA) per milliliter. After standing in an ice bath for approximately 30 minutes with occasional rehomogenization, they were centrifuged for 10 minutes at 2500 g in an International refrigerated centrifuge, model PR-2, to remove debris. Homogenates were kept in an ice bath until used. Dilutions of the homogenate were prepared using the BSA solution as a diluent, the BSA being present to protect the enzyme activity at higher dilutions. The jelly layers of unfertilized eggs and prehatching embryos were removed manually with fine jewelers’ forceps before homogenization. Posthatching larvae and adult tissues were homogenized directly. Wet weights of tadpoles and adult tissues were recorded, and samples of the homogenates were retained for protein determinations as described below. For early developmental stages, ten embryos were homogenized per milliliter of the BSA solution. In later stages the concentration was dictated by the amount of enzyme activity present. Protein
Determinations
Protein was determined by the method of Lowry et al. (1951) modified to permit assay of a 10-J sample of homogenate. A standard curve consisting of several concentrations of BSA was run with each set of determinations. Enzyme
Assays
The assay used for both DNase I and DNase II depends upon the insolubility of macromolecular DNA in cold 2% perchloric acid (PCA). Ten milliliters of appropriate substrate solution was placed in 25-ml Erlenmeyer flasks fitted with ground-glass stoppers, clamped to a Burrell wrist-action shaker, and oscillated gently in a constanttemperature water bath at 27°C. To start the reaction 2 ml of homogenate was added; at 0, 30, 60, and 120 minutes, l-ml aliquots were pipetted in duplicate into chilled tubes, each containing 0.2 ml of cold 12% PCA, mixed thoroughly, and allowed to stand in an ice bath for 15 minutes. The precipitate was sedimented by centrifugation at 2000 g for 10 minutes in an International refrigerated centrifuge. The optical density of the supernatant fluid, containing oligonucleotides
DNASE
ACTIVITIES
IN
FROG
DEVELOPMENT
235
resulting from the degradation of DNA, was read at 260 rnp on the Beckman DU spectrophotometer against an appropriate blank. Ultraviolet absorption spectra of the PCA supernatants were determined on the Beckman DK-2 recording spectrophotometer. One unit of DNase activity is defined as that amount of activity which causes an increase in absorbancy at 260 rnp of 0.001 per 30 minutes of incubation at 27°C. The pH of each flask was determined at the end of each experiment. Substrate solutions were made up fresh every 2 or 3 days, and the DNA stock solution (2 mg DNA per milliliter in distilled water) approximately every week. Validity of the method was initially ascertained by using partially purified commercial DNase (Worthington Biochemical Corporation). Highly purified salmon sperm DNA (California Corporation for Biochemical Research) was used as substrate throughout. Buffers were acetic acid-sodium acetate, Tris-hydrochloric acid, or glycine-sodium hydroxide, as specified in the text. To study the development of DNase activities, approximately equal numbers of eggs were taken from each of four gravid females, fertilized with the same sperm suspension, and sampled at daily intervals until Shumway stage 25 was attained. Determinations were made at approximately the same time every day. Reaction flasks were set up for each enzyme determination as follows: 1. 2. 3. 4.
Homogenate plus buffer BSA plus substrate Homogenate plus substrate Homogenate plus substrate plus inhibitor DNase I inhibited by 0.01 M EDTA DNase II inhibited by 0.05 M MgSO,
Because this investigation involved the use of crude homogenates of developing embryos in which unknown phosphodiesterase activities might be present, it was considered desirable to include a more direct control than that provided by summation of flasks nos. 1 and 2. Thus flask no. 4 was designed to measure the maximal amount of hydrolytic activity that could occur when the activity of DNase I or II was inhibited. When it was expedient to do so, flask no. 4 was set up as a duplicate of flask no. 3. This invariably led to very close duplication of results.
236
JOHN
R. COLEMAN
RESULTS
Before investigating the development of DNase activities in ontogenesis, it was necessary to attempt to define conditions that would be optimal for enzyme assay. DNase
I
Homogenates of tadpoles beyond frog pancreas were found to contain
Shumway stage 25 and adult a DNase activity that required
4000 A
Frog
pancreas
0
Frog
liver
DNase
DNase
6.0
7.0
I
1I
z w
3000
k cc a F L a? u)
2000
s 0 ?
IO00
z 3
1
5.0
0.0
9.0
10.0
PH FIG. 1. Effect of pH on DNase activities of adult frog tissue homogenates. Composition of substrate for DNase I: pH 5.0-6.0, 0.025 M acetate buffer; pH 7.0-9.0, 0.025 M Tris-HCl buffer; pH 9.5, 0.025 M glycine-NaOH buffer; all contained 08.025 M MgS04 and 0.2 mg DNA per milliliter. Composition of substrate for DNase II: pH 4.0-6.3, 0.1 M acetate buffer; pH 7.0-9.0, 0.1 M TrisHCl buffer; all contained 0.1 M NaCI, 0.001 M EDTA, and 0.2 mg DNA per milliliter. Concentration of homogenate for DNase I was 0.07 mg protein per milliliter, for DNase II was 0.14 mg protein per milliliter.
DNASE
ACTIVITIES
IN
FROG
237
DEVELOPMENT
400 DNose o DNose
A
I n
300 -i \ W
cn
I
0
0.1
0.2
mg PROTEIN
0.3
/ml
0.4
0.5
0.6
0.7
HOMOGENATE
FIG. 2. Effect of homogenate dilution on tadpole DNase activities. Composition of substrate for DNase I: 0.025 M Tris-HCl, pH 8.6, 0.005 M MgSO,, 0.2 mg DNA per milliliter; for DNase II: 0.3 M acetate buffer, pH 4.8, 0.001 M EDTA, 0.2 mg DNA per milliliter. DNase I curve is from a whole 36-mm tadpole, Taylor and Kollros stage 4; DNase II curve is from a Taylor and Kollros stage 13 tadpole tail.
magnesium and exhibited optimal activity at pH 8.59.0 in 0.025 M Tris-HCI buffer (Fig. 1) . This activity was designated DNase I. At a DNA concentration of 0.2 mg per milliliter, the optimal MgSO, concentration was found to be in the range of 0.0054025 M, varying slightly with the preparation. Concentrations of MgSO, above 0.025 M were increasingly inhibitory. Omitting MgSO, from the substrate medium reduced the activity to about 20% of that found in the presence of 0.005 M MgSO,. That this requirement is for divalent cations was indicated by the finding that a chelating agent, ethylenediamine tetraacetate (EDTA ) , was capable of inhibiting the reaction. As
238
JOHN
R. COLEMAN
little as 0.0005 M EDTA reduced the activity to about 2% of that found in the presence of magnesium. This inhibition could be overcome by increasing the concentration of MgSO,. When the specific activity of tadpole DNase I was plotted as a function of the DNA concentration in 0.025 M MgSO*, saturation with substrate was found at a concentration of 0.2 mg DNA per milliliter. Therefore, at this substrate concentration, the amount of activity observed during the 2-hour incubation should be directly proportional to the concentration of enzyme. Determinations of the DNase I activity in various dilutions of the homogenate showed that this was the case (Fig. 2).
WAVELENGTH
mp
FIG. 3. Absorption spectra of PCA supernatant digestion of DNA by tadpole DNase I. Composition HCl buffer, pH 8.7, 0.005 M MgSO,, 0.2 mg DNA of homogenate was 0.16 mg protein per milliliter. stage 25.
solutions during course of of substrate: 0.025 M Trisper milliliter. Concentration Tadpoles were Shumway
controls in Ultraviolet absorption spectra, as well as appropriate which DNA or homogenate was omitted from the reaction mixture, indicated that the reaction observed with this assay technique was breakdown of DNA following its exposure to an appropriate homogenate (Fig. 3). Table 1 shows the substrate composition which was found to be optimal for the assay of frog DNase I activity.
DNASE
ACTIVITIES
IN
TABLE
FROG
239
DEVELOPMENT
1
COMPOSITION OF SUBSTRATES FOR DEVELOPMENTAL STUXES DNase I
DNase II
0. u25 &I Tris-HCl 0.005 XI MgS04 0.2 mg DNA per ml pH 8.7 2~ 0.1 p = 0.03
U. 3 AI acetate 0.001 M EDTh 0.2 mg DNA per ml pH 4.9 x!z 0.1 p = 0.18
DNase II Another DNase, designated DNase II, was found in adult tissue and tadpole homogenates. This activity exhibited an optimum at pH 4.5-5.0 at all stages tested (Shumway stage 19 embryo, 3-month-old tadpole, adult frog liver) (Fig. 1). DNase II does not require divalent ions, but is inhibited by them. Thus, a concentration of MgSO, of 0.01 M inhibited the activity 95%. On the other hand, optimal activity was exhibited at an ionic strength ( ,u) of 0.18 in the presence of monovalent ions as provided by acetic acid-Na acetate buffer, or by mixtures of acetate buffer and NaCl. As recommended by Laskowski ( 1961), a small amount of EDTA (0.001 M ) was included in the reaction mixture to prevent inhibition of DNase II activity by divalent cations in the crude homogenates (Table 1). As the concentration of DNA in the reaction mixture was increased, it was found that activity decreased, i.e., substrate inhibition was encountered. At a DNA concentration of 0.2 mg/ml, the homogenate dilution curve was linear (Fig. 2), indicating that the amount of activity observed was proportional to the concentration of enzyme. Substrate inhibition was apparent only in the failure of the curve to bisect the origin. Since substrate became limiting too rapidly at lower DNA concentrations, 0.2 mg DNA per milliliter was selected for use in the developmental studies (Table 1). As in the case of DNase I activity, ultraviolet absorption spectra and appropriate controls indicated that the reaction being followed was the hydrolysis of DNA. For both DNase I and DNase II activities, the increase in absorbancy of the PCA supernatant solution was not a strictly linear function
240
JOHN
R. COLEMAN
of the time of incubation (Fig. 4). The rate of appearance of acidsoluble material absorbing at 260 rnp increased with the time of incubation until the substrate concentration became limiting, after which the rate decreased, resulting in a sigmoid curve. In spite of this, the linearity of the homogenate dilution curves derived from the 60-120 minute interval of incubation (Fig. 2) justified a quanti,600 n
DNose
I
0
DNose
H
: N ti
m u
,400
0
30 TIME
60 IN
90
I20
MINUTES
FIG. 4. Progression of DNA digestion by larval homogenates. of substrates same as for Fig. 2. Concentration of homogenate for 1.1 mg protein per milliliter; for DN ase II, 0.44 mg protein Larvae were early Shumway stage 25.
Composition DNase I was per milliliter.
tative consideration of the amount of nuclease activity present. For technical reasons, the 0 time values were occasionally erratic and initial reaction rates could not be accurately estimated. Consequently, the units of DNase activity reported herein refer to the portion of the curve obtained between 60 and 120 minutes of incubation. Comparable determinations with commercial bovine pancreatic DNase I gave similar results. Dezjelopment
of DNase Activities
During the determination of optimal conditions for enzyme assay, it was found that both DNase activities were low in early embryonic stages, but present in tadpoles and adults. On this basis, it was decided to undertake a rigorous examination of both types of activity
DNASE
ACTIVITIES
IN
FROG
DEVELOPMENT
241
simultaneously in developing embryos until both activities were present in significant amounts. Under the conditions employed, DNase I activity was essentially undetectable prior to stage 23, the beginning of opercular fold formation (Figs. 5 and 6). Activity began to increase slightly at stage 23, 800
700
5
600-
& LL Q.
500-
-
: \
400
-
300
-
200
-
: z cl if z 3
FG
N 18 20
STAGE
22
OF
24
25
Todpolep
DEVELOPMENT
FIG. 5. DNase I activity per milligram protein of the frog. Substrate composition was as shown Development is expressed in Shumway stages.
during early development in Table 1 for DNase
I.
but did not increase rapidly until the operculum was closing at stage 24 or early stage 25. On a protein basis (Fig. 5), the activity appeared to reach a peak and then decline, but the data were quite variable in this region of the curve. On a per embryo basis, this same peak was observed, but activity thereafter continued to increase rather than level off or drop (Fig. 6). From its first appearance at stage 23, the nuclease activity was 90-95% inhibitable by 0.01 M EDTA. A very low level of DNase II activity was found in the unfertilized
242
JOHN
R. COLEMAN
egg, and this dropped slightly through gastrulation (Figs. 7 and 8). With the advent of neurulation, however, this activity began to increase and, on a per milligram protein basis, continued to increase until about stage 24, when it began to level off and perhaps declined
0
F G N 18
STAGE
FIG. 6. Same data
20
22
OF
DNase I activity per embryo as those plotted in Fig. 5, but
24
25
Tadpoles-
DEVELOPMENT
during early development of the frog. calculated on a per embryo basis.
somewhat (Fig. 7). On a per embryo basis, such a change in rate was observed only as a slight shoulder in the curve, after which the activity continued to increase (Fig. 8). The activity observed in these determinations was completely inhibitable by 0.05 M MgSO,. Since the rapid increase in DNase I activity at early stage 25 was approximately coincident with the onset of feeding, it was considered that this activity might be localized in the digestive tract. Stage 25 tadpoles were grown for 1 week to a length of 17 mm, dissected in cold isotonic saline, and the component parts assayed for DNase I activity (Table 2). Gut had some activity, but pancreas was found to be by far the most active tissue examined. DNase II was not com-
F G N
18
20 22
STAGE
24
OF
25
Tadpole-
DEVELOPMENT
FIG. 7. DNase II activity per milligram protein of the frog. Substrate composition was as shown Development is expressed in Shumway stages.
in
during Table
early development 1 for DNase II.
600
F Lx !ii w \
500
400
2 D fi
300
c z 1
200
100
F G N I8
STAGE
20
22
OF
24
25
Tadpoles-
DEVELOPMENT
FIG. 8. DNase II activity per embryo during early development frog. Same data as those plotted in Fig. 7, but calculated on a per embryo Development is expressed in Shumway stages. 243
of
the basis.
244
JOHN
parably localized, although than in the body (Table pancreas.
H. COLEYIAN
its specific activity was higher in the gut 2). In this case, gut included liver and TABLE 2
DISTRIBUTIOK ACTIVITIES
IN
Units Preparation
OF DNASE
IlNnse
Embryo
Whole Gut
SHUMWAY
Pancreas
DNASE
AXD
25
II
TADPOLIW
I *er:
l:nits
DNrtse
II per:
Mg protein
Embryo
406 103 1830 20 !I
689 270
:w1 500
32
350
288 68 330 2 6
Liver Body
I
STAGE
Mg protrin
a Tadpoles were grown to a length of 17 mm and dissected in cold isotonic saline. The digestive tract, with liver and pancreas attached, was separated from the rest of the body after severing the gut immediately anterior t,o the stomach and at the opening into the cloaca. The portion remaining behind is designated “body.” For DNase I, “gut” represents the digestive tract from which liver and pancreas have been removed and assayed separately. For DNase II, “grit” includes liver and pancreas. “Whole” represents undissected tadpoles.
Activities were too low and the embryonic systems were changing too rapidly to permit comparable dissections designed to localize the early increase in DNase II activity. It has been hypothesized that TABLE DNASE
II AND
(Taylor
IICTIVITY
3
TAILS
NONMETAMORPHOSING
stage and liollrou)
13
13 18 2lY 22 n Beginning
IN
Units DNa8c II per mg protein
348 395 705
OF METAMORPHOSIW TAI)POLES l!nitti DKase II per rng wet weight
8.0 8.6 18.3
1170
31.7
1160
il.8
of tail resorption.
this hydrolytic enzyme may play a role in degenerative processes (cf. de Duve, 1959; Novikoff, 1961). To investigate its correlation with a degenerative process in the frog, the DNase II activities of homogenates of tadpole tails were determined before and during resorption
DNASE
ACTIVITlES
IN
FROG
DEVELOPMENT
945
of the tail at metamorphosis. Tails in the process of resorption had severalfold the DNase II activity of tails not undergoing resorption (Table 3). DISCUSSION
The failure to detect DNase I activity in the early frog embryo reported in this work is in apparent conflict with the results of Finamore (1955). However, the activities reported by Finamore were very low, were obtained at suboptimal pH (pH 7.4), and apparently were not controlled for homogenate autolysis. Hence, his results do not necessarily reflect nuclease activity. Blumenthal (1957) was unable to detect DNase I activity in embryos, tadpoles, or adults of RUMZ pipiens. In this case, failure to detect activity was probably due to the use of suboptimal pH (pH 4-7) in combination with high ionic strength (0.2 M phosphate buffer) leading to complete inactivation of DNase I (cf. Rotherham et al., 1956). Blumenthal found DNase II activity in adult frogs, but was unable to detect it in embryos or tadpoles. These results led her to suggest that this enzyme probably appeared at metamorphosis. In the present investigation, DNase II activity was detected at all stages of development, although activity was very low prior to neurulation. In Blumenthal’s work, it is likely that a high substrate concentration (1.5 mg DNA per milliliter) in combination with low ionic strength (0.1 M acetate buffer, ,U approximately 0.06) prevented observation of DNase II activity in any but the most active preparations. The determination of optimal assay conditions permits, as a corollary, comparison of the reaction characteristics of frog DNase I and DNase II activities with the more extensive information available on mammalian DNases (reviewed by Laskowski, 1961). Except for the pH optimum of frog DNase I activity, other characteristics of the DNase activities reported here are quite similar to those of the DNase I and DNase II activities studied in mammals. In fact, the conditions for the assay of frog DNase II activity derived from the investigations reported here are very close to those recommended by Laskowski (1961) for the study of DNase II in higher vertebrate tissues. The exceptionally alkaline pH optimum of frog DNase I activity may represent a real divergence from the type of DNase I activity found in higher forms or may be a function of the use of crude homogenates. Siebert et al. (1950) reported considerable variation in the pH opti-
mum for DNase activity in crude homogenates depending on the tissue and animal investigated. Substrate inhibition is commonly encountered in studies of both DNase I and DNase II (0th et al., 1958; Laskowski, 1961) although, in the present investigation, it was encountered only with DNase II activity. The extent to which this may reflect the presence of divalent cations in the DNA preparation is not clear, but 0th et al. report no change in activity after dialysis of DNA against water or EDTA. The linearity of the homogenate dilution curves (Fig. 2) justifies the consideration of DNase activities in quantitative terms in spite of substrate inhibition and the nonlinear rate of appearance of reaction products (Fig. 4). The sigmoid relationship of the appearance of acid-soluble products with time of incubation is intimately related to the endonuclease nature of these enzymes, rather than being a consequence of the use of suboptimal conditions and crude homogenates. Similarly complex characteristics seem to prevail even in studies of purified DNase I (Kunitz, 1950) and DNase II (see Laskowski, 1961, for detailed discussion). Comparison of DNase I and DNase II activities in the frog embryo shows that they undergo strikingly different patterns of development (Figs. 5-B). The initial rise in DNase II activity approximately coincides with neurulation, occurring much earlier and at a slower rate than the DNase I activity. DNase I begins to increase during at which time the yolk reserves are beopercular fold formation, coming depleted, and approaches maximal activity at about the time active feeding begins at Shumway stage 25. On a protein basis, the increase in DNase II activity ceases at about the time DNase I activity becomes detectable (Figs. 5 and 7). The development of DNase I activity between stages 23 and 25 can be correlated rather closely with the morphological development of the amphibian pancreas as reported by other investigators (Huettner, 1949; Rugh, 1951; Tahara and Nakamura, 1961) and, in fact, this activity may be localized in the pancreas (Table 2). In investigations on mammals, the detection of DNase I activity in tissues other than pancreas is frequently complicated by inhibitors (Dabowska et al., 1949; Cunningham and Laskowski, 1953; Feinstein and Hagen, 1959). The relatively high activity of the isolated pancreas compared with the activity of the whole tadpole (Table 2) may reflect
DNASE
ACTIVITIES
IN
FROG
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the presence of such inhibitors in tadpole tissues, although no critical attempts to demonstrate them have been made. The correlation between the development of DNase I activity and the onset of active feeding at stage 25 lends support to the hypothesis of Allfrey and Mirsky (1952) that the function of pancreatic DNase I is primarily digestive. The DNase I activity found in gut homogenates (Table 2) may thus be of pancreatic origin. Relatively few of the enzymes studied in anuran embryos increase as late in embryonic development as does DNase I activity. Those which do include a-amylase (Scollo-Lavizzari, 1956), an alkaline protease, and lipase in Bufo culgaris (Urbani, 1957), an alkaline protease in Xenopus (Lovtrup, 1955), and DPNH-cytochrome c reductase in Rana pipiens (Lang and Grant, 1961; Wallace, 1961). No attempts at tissue localization have been reported for these enzymes, but at least the first four mentioned could reflect pancreatic development. In contrast with DNase I, DNase II activity began to increase during neurulation; at this time localized degenerative processes have been reported to occur in the formation of the neural plate and neural tube (Glucksmann, 1951). In mammals, DNase II has been shown to be compartmentalized with other acid hydrolases in the lysosomal fraction of tissue homogenates (de Duve et al., 1955; van Lancker and Holtzer, 1959) and has been hypothesized to play a degradative role in regions of cellular degeneration (see de Duve, 1959, and Novikoff, 1961, for extensive reviews). The increase in DNase II activity in the early neurula may reflect such degenerative processes, but for technical reasons no attempt was made to determine the regional distribution of DNase II activity in the early embryo. By Shumway stage 25, the activity appears to be widespread (Table 2). Of other acid hydrolases studied in anuran embryos, acid protease ( Urbani, 1957)) acid phosphatase and acid RNase (de Cesaris Coromaldi, 1955, 1958) begin to increase in the tailbud stage of Bufo uulgaris, whereas cathepsin increases in the gastrula stage of Xenopus Eaewis (Deuchar, 1958). Information concerning the extent to which these developmental differences might reflect differences in compartmentalization is currently lacking. An .attempt was made to correlate the activity of DNase II with a more accessible degenerative process, the resorption of the tadpole
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tail at metamorphosis. On a protein basis, tails in the process of resorption were found to have about three times the DNase II activity of nonresorbing tails (Table 3). On a wet-weight basis, this figure is enlarged to fivefold. These results are in close agreement with those of other workers on hydrolytic enzymes in anuran metamorphosis (Urbani, 1957; Weber, 1957; de Cesaris Coromaldi, 1958; Weber and Niehus, 1961) and have been confirmed in more extensive investigations of thyroxine-induced tail resorption in R. catesbeiana (Coleman, unpublished). Finally, no critical information is available concerning the extent to which the increases in nuclease activities reported in this paper reflect de novo synthesis of new enzyme protein. Preliminary experiments with mixtures of homogenates from different stages of development indicate that inhibitors and/or activators may not play a major role in the regulation of the observed DNase activities. Since DNase II activity increases at a time when the embryo depends completely on its stored yolk reserves, more sophisticated biochemical approaches are difficult. On the other hand, the relatively late increase in DNase I activity may permit investigation of this question through the use of radioisotopes or immunological cross reactivity, pending preliminary purification of the enzyme proteins involved. Until the problem of de now synthesis is clarified, little can be said about the mechanisms that regulate enzyme activity or synthesis. SUMMARY
Deoxyribonucleases have been studied in the leopard frog, Rana pipiens, by following the progressive increase of perchloric acidsoluble compounds absorbing at 260 rnp during incubation of homogenates with macromolecular DNA at 27°C. The terms DNase I and DNase II, as used in this report, do not necessarily imply identity with purified DNase I and DNase II as studied in mammals, but refer only to the enzymatic breakdown of DNA under two given sets of conditions. Frog DNase I activity exhibits an alkaline pH optimum of pH 8.5-9.0, but in other requirements is similar to DNase I activities studied in higher vertebrates. It is relatively unaffected by excess substrate, exhibits a divalent cation requirement which can be satisfied by magnesium but is inhibited by concentrations of MgSO, above 0,025 M, Frog DNase II activity is similar to mammalian DNase II
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in all the requirements studied. It has a pH optimum of 4.5-5.0, exhibits a high degree of sensitivity to excess substrate, requires an ionic strength of about 0.18 for optimal activity, and is strongly inhibited by divalent ions. DNase I activity was found in adult pancreas and whole tadpole homogenates. By the assay technique employed, it was undetectable in the early embryo but began to rise at approximately Shumway stage 23. This coincides approximately with the time of morphological appearance of the anuran pancreas as reported in the literature. Stage 25 tadpoles grown for 1 week and dissected showed a striking localization of DNase I activity in the pancreas, and perhaps some in the gut as well. Low DNase II activity was found in the unfertilized egg. This activity decreased slightly through gastrulation, then began to increase with the onset of neurulation. On a per milligram protein basis, this activity continued to increase through early prelarval development but leveled off at about the time DNase I activity became detectable. On a per embryo basis, the activity continued to increase. Hence, these two nuclease activities show strikingly different patterns of development. The increase in DNase II activity at neurulation coincides with cellular degeneration reported to occur in the forming neural plate and neural tube of vertebrate embryos. No attempt was made to determine the localization of DNase II activity in the neurula. A three- to fivefold increase in activity was found during tail resorption at metamorphosis, a result which suggests a correlation between DNase II activity and degenerative processes. The author gratefully acknowledges the persistent encouragement tion of Dr. James D. Ebert, Dr. Yoshihiro Kato, and Dr. Annette Acknowledgment is also extended to Dr. Heinz Herrmann for his of the manuscript.
and
stimulaColeman. critical reading W.
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