Biochimica et Biophysics Elsevier
BBALIP
19
Acta, 1046 (1990) 19-26
53458
Purification and partial characterization of intestinal acid lipase R. Hanumantha
Rao and Charles M. Mansbach,
II
Department of Medicine, Division of Gastroenterology, The University of Tennessee, Memphis and The Veterans Administration Medical Center, Memphis, TN (U.S.A.)
(Revised
Key words:
Lipase;
Intestine;
(Received 23 January 1989) manuscript received 18 April 1990)
Lipolysis;
Protein purification;
Intestinal
mucosa;
Lipid metabolism
Intestinal acid lipase is an enzyme whose greatest specific activity is localized to the villus tips of the proximal intestine (Rae, R.H. and Mansbach, C.M. (1990) Biochim. Biophys. Acta 1043,273-280). This suggests that it plays a role in the processing of dietary lipids. We purified the enzyme in order to better characterize it. Acid lipase was isolated from intestinal mucosa of rats by a combination of ammonium sulfate precipitation, butanol extraction and chromatography on DEAE Bio-Gel, CM Bio-Gel and Sephadex G-75. This resulted in a single protein of M, 53700 on SDS-polyacrylamide gel electrophoresis. The isolation scheme produced a 3344-fold purification resulting in an enzyme whose specific activity was 801 pmol/min per mg protein. The yield was 50%. The purified enzyme was stimulated @&fold) by the addition of tauro- or glycocholate but no other conjugated bile acid. A sharp peak in activity occurred at pH 5.6. The pl of the enzyme was 6.2. The reaction products produced under prolonged incubation suggested that monoacylglycerol was not hydrolyzed since an overabundance of monoacylglycerol was found with respect to the amount of fatty acid produced. These results suggested that intestinal acid lipase is potentially important in the metabolism of dietary lipids. Its proportionate role awaits further documentation.
Introduction A number of acid active Iipases, usually lysosomal in origin, have been purified from various tissues and species. The activity of these Iipases varies from pmol to nmol/mg protein per min. Those lipases with the greatest activity are secreted into the gastrointestinal tract. These include the pharyngeal [l], lingual [2] and gastric lipases [3,4]. Those with the least activity are the ones isolated from tissues such as the fibroblast [4] which normally do not process much lipid. We have recently studied a similar lipase of intestinal origin [5]. This hpase appeared to be in position to play a role in the intestinal metabolism of absorbed lipids based on the circumstantial evidence that it is found in greatest activity in the proximal intestine in the villus tips. This is the location expected for enzymes and transporters engaged in the absorption and processing of dietary substrates. The quantitative role of this enzyme in lipid metabolism remains to be explored, however.
Correspondence: see, Memphis, 38163, U.S.A.
Charles M. Mansbach, II, The University of Tennes951 Court Avenue, Room 555 Dobbs, Memphis, TN
0005-2760/90/$03.50
0 1990 Elsevier Science Publishers
B.V. (Biomedical
The current investigations were conducted toward purifying acid lipase from the intestine. Previously purified acid Iipases [l-4,6-18] have shown remarkable similarity with regard to their pH optima, their molecular weight and their pl, but not their activity against long acyl chain (C > 12) triacylglycerol (TG) substrate. We wished to determine if the intestinal enzyme were similar to those previously reported with respect to these physicochemical characteristics. Equally important, with respect to considerations for a physiological role of the enzyme, is the specific activity of the purified enzyme. If high, it would more likely play a role in intestinal lipid metabolism. Methods Tissue preparation Male Sprague-Dawley rats weighing 250-350 g were used. Usually 16 rats were processed at a time. Under ether anesthesia, their intestines were removed to an iced glass plate. The mucosa from the proximal 3/4 intestine was harvested by the aid of giass microscope slides and placed in Buffer A (taurocholate, 4 mM; dithiothreitol (DTT), 2.5 mM; Tris-HCl, 10 mM (pH 7.4). The mucosa was homogenized in 10 wt/v Buffer A using a gIass homogenizer with Teflon pestle. The hoDivision)
20 mogenate was centrifuged for 3. lo5 g. min in a Sorval RC 5C centrifuge (DuPont Instruments, Norwalk, CT). The pellet was extracted three times with 50 ml of Buffer A and re-centrifuged in between. The supernatants were combined and adjusted to 50% saturated (NH4)*S04 and stirred gently at 2°C for 30 min. The mixture was then centrifuged for 3.10’ g. min. The resulting pellet was resuspended in Buffer A supplemented with 20% ethylene glycol (Buffer B) and dialyzed overnight with two changes of buffer. The dialysate was extracted with butanol at 2°C which was added slowly until it reached 0.5 volumes. The mixture was stirred for 20 min and then centrifuged for 8. lo4 g. min. The butanol was removed by suction and the lower, aqueous layer re-extracted with butanol as previously. The proteinaceous material that forms at the butanol/H,O interface was discarded. The combined aqueous phases was dialyzed against two changes of buffer B. Chromatographic procedures The dialysate was applied to a DEAE Bio-Gel A (100-200 mesh, Bio-Rad, Richmond, CA) column (1.5 x 24 cm) without prior concentration. The column was pre-equilibrated with Buffer B and eluted with the same buffer. Active fractions were concentrated on a YM 10 membrane (Amicon Corp, Danvers, MA) and dialyzed against Buffer C (sodium acetate, 20 mM; dithiothreitol (DTT), 2.5 mM; ethylene glycol, 20% and taurocholate, 4 mM; pH 5.5). The buffer was changed three times. The dialysate was placed on a CM Bio-Gel (Bio-Rad Laboratories, 50-200 mesh) column of 1.5 x 24 cm. The column was pre-equilibrated with Buffer C and eluted with a NaCl gradient of 0 to 0.4 M using a gradient mixer. The flow rate was 4.8 ml/h. 2-ml fractions were collected. The active fractions were dialyzed against Buffer B with three changes of dialysand and concentrated on a YM 10 membrane (Amicon Corp). The concentrated material was applied to a Sephadex G-75 superfine column (Pharmacia Fine Chemicals, Piscataway, NJ) of 2.5 X 65 cm and eluted with Buffer B. The column flow rate was 6 ml/h. 2-ml fractions were collected. The column was calibrated using bovine serum albumin, 66 kDa; ovalbumin, 45 kDa; chymotrypsinogen, 25 kDa; and ribonuclease A, 13.7 kDa (all supplied by Sigma Chemical, St. Louis, MO). All chromatographic procedures were performed in a cold room. SDS polyacrylamide gel electrophoresis (SDS PAGE) was performed in a Hoefer model 500 apparatus (Hoefer Scientific Instruments, San Francisco, CA) as suggested by Laemmli [19]. Standard proteins of known M, (see legend to Fig. 3) or samples of acid lipase were heated to 100°C for 4 min in the presence of SDS, 2% and /3-mercaptoethanol, 5%. 20 pg of acid lipase was applied to the 10% cross-linked gel and electrophoresed for 5 h using approx. 30 mA per gel. After development, the gel
was stained with 0.1% Coomassie Blue R-250 in 40% methanol and 10% acetic acid. It was destained with 10% methanol and 7.5% acetic acid. Isoelectric focusing was performed using a LKB 110 ml column (LKB Washington, DC) cooled to 2” C. The ampholyte used was Pharmalyte (Pharmacia, Uppsala, Sweden) which was used at a final concentration of 2% at pH 4.0-6.5. A gradient of O-40% glycerol was constructed and the enzyme applied to the middle of the column in 0% glycerol. The protein used was the dialyzed soluble extract (see Table I). The total run was 18 h at 1600 V. Protein modifying reagents The purified enzyme was preincubated for 20 min in the presence of inhibitor at 37” C. Acid lipase activity remaining was assayed for 2 min as in Analytical Methods. The following potential inhibitors were utilized (Sigma Chemical): N-ethylmaleimide, N-bromosuccinimide, 2,4,6-trinitrobenzenesulfonic acid, diethyl pyrocarbonate and diethyl p-nitrophenyl phosphate (E600). Substrate specificity studies Purified enzyme was incubated in the presence of either glyceryl tri[i4C]oleate, l-palmitoyl-2-[‘4C]oleoylphosphatidylcholine (New England Nuclear), cholesteryl [i4C]oleate (New England Nuclear), 4-methylumbelliferyl oleate or, 4-methylumbelliferyl palmitate (both from Sigma Chemical) in the taurocholate containing buffer described in Analytical Methods. The incubations were for 2 min each at 3 mM substrate. Released oleate from TO and cholesteryl oleate were quantified as in Analytical Methods. Potential phospholipase A, or A, activity was determined by extracting the incubation mixture [20] and separating the products on TLC using as the mobile phase chloroform/ methanol/ acetic acid/H,0 (25 : 15 : 4.2, v/v) [21]. Lyso derivatives and fatty acids, the potential products, were identified by authentic standards. The appropriate areas were scraped from the plate and radioactivity determined (Analytical Methods). The production of umbelliferone was quantified by fluorometry [6] with a Hitachi model 650-15 fluorecence spectrophotometer (Hitachi, Tokyo, Japan) using 320 nm as the excitation wavelength and 450 nm as the emission wavelength. Analytical methods Acid lipase was assayed using glyceryl tri[14C]oleate (TO) (New England Nuclear, Boston, MA), 3 mM (1 . lo5 dpm/pmol); sodium taurocholate, 20 mM (Calbiochem, La Jolla, CA); DTT, 1 mM; glycerol, 0.8 M; sodium acetate, 50 mM (pH 5.8) and appropriate amounts of protein depending on the degree of purity in a total volume of 250 ~1 [5]. The TO was prepared for assay by sonifying it in 4% glycerol and 4 mM tauro-
21 cholate. The reaction was stopped and released oleate extracted by the addition of 0.5 ml carbon tetrachloride/ hexane (2 : 1, v/v) containing 1 mM Triton X-100 and 0.1 mM oleate, and 0.75 ml of ethanol/ water (3 : 1, v/v) containing 70 mM NaOH [6]. Radioactivity was determined using Scintiverse II (Fisher Scientific) in a Packard 1500 liquid spectrometer (Packard Instruments, Downers Grove, IL). 96% of the extracted radioactivity was FFA. The Folch [20] extraction procedure was tested for its ability to extract the total FFA present by extracting TO (750 nmol) plus a known amount of [‘4C]oleate (250 nmol). 98.8% of the dpm were in the organic phase. 85% of lipid dpm after a lipase incubation that were subjected to TLC were recovered from the plate as compared to the direct determination of radioactivity. Protein was determined by the method of Lowry et al. [22]. Thin-layer chromatography (TLC) was performed on Silica-gel H layers using the solvent system hexane/ diethyl ether/ methanol/ acetic acid (80 : 20 : 6 : 2, v/v).
Results The steps used in the purification of intestinal acid lipase are shown in Table 1. It is evident that solubilization of the enzyme into taurocholate micelles enhanced its activity as compared to that present in the whole homogenate. This resulted in more enzymatic activity than was originally present in the non-processed homogenate. This may have been due to the solubilized enzyme having a preferred conformation for interaction with substrate. Alternatively, an inhibitor could have been present which pelleted on centrifugation. Multiple
other detergents were tried in attempts to solubilize the enzyme as previously reported [5]. All of these inhibited enzyme activity [5]. A O-50% ammonium sulfate precipitation step was used to concentrate the enzyme and to remove soluble proteins. This resulted in a 3.8-fold purification (Table I) which is at the upper end of the expected range (2-4-fold). The resulting material had moderate turbidity and could not be loaded onto chromatographic columns. In order to more completely solubilize the enzyme and to remove associated membranous fragments, a butanol extraction step was employed. 0.5 vol. of butanol (2” C) was added slowly while stirring which was continued for 20 min. The mixture was centrifuged for 38 000 x g. min to separate the phases and the butanol layer removed by suction and discarded. The cake which formed at the butanol/aqueous interface contained 24% of the enzymatic activity. The interfacial proteins were extracted with Buffer A. Only 10% of the activity could be recovered. Therefore, the interfacial proteins were discarded as well. The aqueous phase was extracted a second time with butanol and the phases separated as before. The aqueous phases were combined. The combined aqueous phases had an increase in specific activity of only 1.6-fold (Table I) as a result of the butanol treatment. This step was necessary, however, to yield a clear solution which could be chromatographed. The aqueous phase from the butanol extract was applied to a DEAE Bio-Gel column (1.5 X 24 cm) after dialysis against buffer A. It was eluted with the same buffer. The enzyme did not bind to the column. Nevertheless, considerable unwanted proteins did which resulted in a 6.4-fold (Table I) purification. Since no binding occurred, the data are not presented further.
TABLE I Purification of intestinal acid lipase Fraction
Total protein
(mg)
Specific activity ( pmol OA/min per mg protein”)
Total activity ( pmol OA/min)
Recovery b (W)
Purification (fold)
Whole homogenate
2820
0.240
616
100
1
Soluble extract
1585
0.611
968
143
2.5
155
9.4
Ammonium
sulfate PPtn’
Butanol extraction DEAE Bio-Gel
463
2.26
1047
234
3.36
785
116
14.0
827
122
88
39.3
21.0
CM B&Gel
2.30
206
474
70
861
Sephadex G-75
0.418
801
335
50
3344
a Oleic acid rekased. b Percentage recovery is based on that present in the whole homogenate. ’ Precipitation.
22 CHROMATOGRAPHY
OF ACID
CHROMATOGRAPHY
LIPASE ON CM SIOGEL
LIPASE
12
= _E
3
mm*-
44
I_
OF ACID
ON SEPHADEX
G-75
PROTEIN
54
64
?b
0
FRACTION FRACTION
NUMBER
Fig. 1. The proteins which passed through the DEAE Bio-Gel column without adhering to the support were chromatographed on CM BioGel. Acid lipase activity (solid tine) was eluted from the column in response to an NaCl gradient as indicated by the open chain line. Proteins, measured at A,,, are indicated by the dashed line.
The active fractions were concentrated on a YM 10 membrane and dialyzed against buffer B. The enzyme was then fractionated on CM Bio-Gel (1.5 x 24 cm) previously equilibrated with the same buffer. 2-ml fractions were collected. After 30 ml of buffer had passed through the column, a NaCl gradient elution was established with a maximal concentration of 0.4 M. As shown in Fig. 1, at 0.05 M NaCl the enzyme began to elute from the column. Fractions 61-74 were pooled, dialyzed against buffer A and concentrated on a YM 10 membrane. A 9.8-fold purification was obtained (Table I>* The concentrated sample was next separated on Sephadex G-75 superfine using a column 2.5 X 65 cm. The enzyme was eluted with buffer A as a single peak coincident with the protein peak as shown in Fig. 2. Calibration of the column with proteins of known molecular weight suggested a hydrodynamic radius consistent with a M, of 44000. Fractions 67-77 were pooled and concentrated on a YM 10 membrane. A further 4-fold purification was achieved (Table I). In sum, the six purification steps gave a 3344fold purification with a 50% yield based on activity present in the original homogenate. SDS-PAGE was performed on the concentrated G-75 eluent. The results are shown in Fig. 3. By Coomassie blue stain, only one band was visible. No other bands were visualized with silver staining (data not shown). When compared to other proteins of known molecular weight which were similarly electrophoresed, acid lipase had a suggested M, of 53700. It is not clear why the
NUMBER
Fig. 2. Fractions 61-74 from the CM Bio-Gel column were concentrated and passed over a Sephadex G-75 column. Acid lipase activity (solid line) eluted as a single peak with a trailing shoulder. Proteins, measured at A,, and shown by the dashed line, peaked in a coincident manner with the major protein peak. The column was calibrated by chromatographing proteins of known molecular weight. These were: bovine serum albumin (M, 66000), ovalbumin (M, 45000). chymotrypsinogen (M, 25000) and ribonuclease A (M, 13 700). The molecular weight of acid lipase was calculated by plotting the K,, [37] of the known proteins against their molecular weights.
66 KD 45 KD 36 KD 29 KD 24 KD
Fig. 3. Fractions 66-77 from the Sephadex column were concentrated and subjected to polyacrylamide gel electrophoresis in the presence of SDS. The protein migrated as a single band of M, 53700. This was calculated by plotting the R, values of the known proteins against their molecular weights as a semilog plot. These proteins were: bovine serum albumin (M, 66000), ovalbumin (M, 45000), glyceraldehyde3-phosphate dehydrogenase (M, 36000), carbonic anhydrase (M, 29000) and trypsinogen (M, 24000).
23
ISOELECTRIC
FOCUSING
OF ACID
LIPASE
TABLE
II
Effect of various bile salts on intestinal acid lipase activity =
14-
E
.
-.-.‘D--
OD260
+
ACTIVITY
Activity OA b released (nmol/min) ’
Bile salt a
No addition
2.6f0.3
Taurocholate
48.7 f 4.1 3.5 *0.4
Glycodeoxycholate
50.7 + 4.7
Glycocholate
4
2
6
10
6
PH Fig. 4. Isoelectric focusing of acid lipase is shown in the figure. Acid lipase activity is shown by the solid line and protein, as measured at GD,se, by the dashed line. Soluble extract (Table I) was used as the protein source. A pH gradient of 4.0-6.5 was used in the presence of a glycerol gradient (O-401).
enzyme appears to have a molecular weight which is smaller on gel permeation chromatography than when it is chromatographed as the denatured, reduced protein. This has been observed previously with calf pharyngeal lipase [l]. One possibility is an interaction of the enzyme with the gel. This could lead to a spuriously
EFFECT
OF pH ON ACID
LIPASE
ACTIVITY
600-
Glycochenodeoxycholate
6.5 f 0.9
Taurodeoxycholate
6.9kO.8
Taurochenodeoxycholate
6.1 f 0.8
a All bile acids were present at a concentration of 20 mM. b Oleic acid released. ’ The data are the meanfS.E. of four experiments. In each case, blank tubes containing the appropriate bile acid were run in parallel with the tubes containing enzyme.
delayed appearance of the enzyme in the chromatogram which would yield a result suggesting a smaller protein than what was actually present. Isoelectric focusing of the enzyme was performed and the results shown in Fig. 4. A p1 of 6.2 is suggested. The activity yield from this procedure was low so that it was not useful as part of the purification scheme. Acid lipase activity as a function of pH is reported in Fig. 5. A sharp rise in activity is seen between pH 5.2 and 5.6 which is the pH at which maximum activity of the purified enzyme occurs. Enzymatic activity progressively fell at more alkaline pH but persisted into the alkaline range. The activity at pH 8 was, however, 7.5fold less than at pH 5.6.
WO-
TABLE
III
400-
Identification
of reaction products of intestinal acid lipase
soo-
TG
DG
MG
2uo-
Control (no enzyme)
57630
1033
100 -
Experimental a (+ enzyme)
14983
11756
Net change 4.0
5.0
6.0
7.0
6.0
PH Fig. 5. Acid enzyme was 4.0-5.6 and buffers were 5.8) normally
lipase activity as a function of pH is shown. Purified used. Acetate buffer, 0.05 M, was used between pH Tris-maleate buffer, 0.05 M, between pH 5.6-8.4. These used as a substitute for the sodium acetate, 0.05 M (pH present in the assay. The other components of the assay remained the same (see Methods).
Ratio
- 42 647
FFA
477
13167
+10723
18179
+12690 1
2 322
+ 15 857 1.2
1.5
a 0.51 pg pure enzyme was incubated with 750 nmol [‘?I TO (spec. act. 99.8 dpm/nmol, 20 mM taurocholate and 50 mM sodium acetate buffer (pH 5.8) in a total volume of 0.25 ml. The incubation was for 1 h at 37OC in a shaking water bath. The reaction was stopped by the addition of chloroform/methanol (2: 1, v/v) and the lipids extracted [20]. Control incubations were the same except no enzyme was added. The data are the mean of two separate experiments performed in duplicate. Values are in dpm.
24 TABLE
IV
Substrate
a
Glycerol
Fatty acid released b (nmol/min per mg protein)
[i4C]oleate
692
l-Palmitoyl-2-[14C]oleoylphosphatidylcholine
0
Cholesteryl
0
[‘4C]oleate
4-Methylumbelliferyl
oleate
4-Methylumbelliferyl
palmitate
12519 1824
a Each substrate was present in a concentration of 3 mM. b Acid lipase activity was determined as in Materials and Methods. All assays were performed twice in duplicate.
Of particular interest with respect to a mucosal enzyme is the effect of bile salts on enzymatic activity. The purified enzyme was significantly stimulated by trihydroxylated bile salts as shown in Table II. Both glyco- and taurocholyl conjugates were equally effective in increasing enzyme activity. These bile acids stimulated enzymatic activity 20-fold. These data differ significantly with that previously reported in impure preparations which are stimulated by a variety of bile acids
PI. In Table III are shown data with regard to the reaction products produced from TG upon prolonged lipolysis. As shown in the table, 74% of the TG originally present was hydrolyzed after 1 h incubation at 37 o C. As was observed with the crude preparation, an excess of MG was observed for the amount of FFA released. For each DG, one FFA and for MG two FFA should be produced. It is immediately apparent that there is less FFA than would be expected for the amount of DG and MG present. Although the reason
TABLE
V
Effect of protein-modifying
reagents on acid lipase activity
Acid lipase was preincubated for at 37OC. Glyceryl trioleate was for 2 min. Acid lipase activity Materials and Methods. Control was 692 pmol acid released/mm performed twice in duplicate.
20 min in the presence of the reagent added and the incubation continued was determined as described under activity with no addition of reagent per mg protein. All assays were
Reagent
mM Reagent
Activity
No addition
_
100
N-Ethylmaleimide
4.0
6
N-Bromosuccinimide
4.0
113
4.0
49
4.0
100
0.1
36
2,4,6-Trinitrobenzenesulfonic Diethyl
pyrocarbonate
Diethyl
p-nitrophenyl
a Percentage
of control
acid
phosphate values.
(E600)
a
for this is not clear, it is evident that the reaction is likely to stop at MG and that complete hydrolysis of TG does not occur. The substrate specificity of the purified lipase was investigated by its incubation with a variety of potential substrates. The results are shown in Table IV which reveal that it is inactive against cholesteryl oleate and phosphatidylcholine. It is, however, even more active against the umbelliferone esters than against TO. This has been noted previously with other lipases [6,13,14]. The results of inhibitor studies are shown in Table V. These demonstrate that there is likely to be a serine at the active site of the enzyme as both N-ethylmaleimide and E-600 were greatly inhibitory. Partial inhibition was observed using large concentrations of trinitrobenzenesulfonic acid indicating that a lysine residue is near but not at the active site. Inhibitors of histidine (diethyl pyrocarbonate) and tryptophane (bromosuccinimide) did not disturb enzymatic activity indicating that these two amino acids are not near L active site. Discussion We have purified a mucosal, acid active, lipase of M, 54 kDa. This lipase is very active, second only in activity to the acid lipase isolated from the rabbit stomach [4] when TO is used as substrate. It is lysosomal in origin and its maximal activity is in the villus tip cells of the proximal intestine [5]. It is distinguished from preduodenal lipase (lingual lipase in the rat [23]) by its continued activity despite esophageal diversion for 48 h [5] and from pancreatic lipase both by its stimulation by taurocholate rather than its inhibition [24] and by its pH optimum [25]. The mucosal acid lipase isolated is remarkably similar to a variety of other acidic lipases isolated from multiple species and tissues [l-4,5-18]. Of the 17 different reports, the average (*SD.) M, is 53300 _t 1100 excluding the human placental enzyme [13]. The pH optimum of the lipases average 4.8 + 0.5. This average is slightly more acidic than the 5.6 of the mucosal enzyme reported here. Nevertheless, there are five lipases whose pH optima either overlap with the intestinal enzyme or are within 0.3 pH units of it [2-4,12,15,18]. Intestinal acid lipase is acidic as suggested by its pl of 6.2. This is comparable to the p1 of the hepatic enzyme [9] and slightly more acidic than the cardiac myosidic and calf pharyngeal enzymes whose pl are 6.3 and 7.0, respectively (17.1). The similarity of the acid lipases suggests that considerable sequence homology will be identified. Despite these apparent similarities, differences between the intestinal enzyme and other acidic lipases are evident. In response to bile acids, the rat lingual enzyme has been shown to have both increased [2] and decreased [16] activity; the human gastric lipase is inhibited by all bile acids [3]. By contrast, the intestinal
25 enzyme is significantly stimulated by trihydroxylated bile acids. In this regard it is similar to sterol ester hydrolase which is specifically stimulated by taurocholate [26] and acid cholesterol esterase which is also stimulated by taurocholate [27]. With regard to the non-ionic detergent, Triton X-100, the liver enzyme is consistently stable [1,7-91 whereas the intestinal enzyme is completely inhibited. The co-addition of taurocholate partially restores activity of the intestinal enzyme in response to Triton treatment, however [5]. In sum, these data would suggest that the intestinal lipase has an interfacial recognition site which is similar to the pancreatic and cholesterol esterases but different from the acid lipases. The active site of the intestinal enzyme is likely to differ from that of the sterol ester hydrolases in that the lipase does not hydrolyze cholesterol esters. Its inability to hydrolyze phosphatidylcholine also distinguishes it from hepatic lipase which does [28]. The degree to which acid lipases hydrolyze a TG substrate also varies. The rat lingual enzyme completely hydrolyzes TG [2,15] whereas the liver enzyme is not able to hydrolyze monoacylglycerol [6,8]. In this regard, the liver and intestinal acid lipases are similar to pancreatic lipase whose inability to completely hydrolyze TG, leaving monoacylglycerol, is well established [29,30]. The purified intestinal acid lipase is a very active enzyme, with a specific activity of 801 pmol/min per mg prot. This is the most active of all reported acidic lipases with the exception of the rabbit gastric lipase [4]. In general the acid lipases can be divided into three groups according to their specific activities. The lipases with the most activity are those which are secreted into the intestinal lumen. In this group are the gastric juice lipases from both human [3] and rabbit [4]. Both of these have activities similar to the intestinal enzyme. The other lipases in this group are the lingual lipases which also demonstrate considerable activity [1,2] with the exception of one isolation [15]. Although the intestinal enzyme is as active as the gastric and lingual ones there is no evidence that it is secreted into the lumen. The intestinal enzyme has a subcellular localization in lysosomes [5]. As such it would make its extracellular secretory potential unlikely. In the second group with intermediate activity is the liver enzyme. Four different isolates demonstrate activity in the range of 2.5 to 7.1 ~mol/min per mg prot. [6-8,111. One isolate, however, is much more active (408 ~mol/min per mg prot.) [9]. The liver enzyme appears to be involved in the hydrolysis of TG stored in the hepatocyte [31]. Finally, there is a third group of lipases whose activity is quite low, in the nanomolar range. These lipases have been isolated from organs that normally process little lipid such as the thyroid [12], fibroblasts [14], placenta [13], cardiac myocytes [17] and leukocytes [18].
The high specific activity of intestinal acid lipase would suggest that it does in fact play a role in the normal intestinal processing of dietary lipid. This potential is enhanced by the finding that in neonates who congenitally lack acid lipase activity in their intestines large lipid droplets are found at electron microscopy [32]. Additional direct evidence comes from experimental studies in rats in which it has been shown that the administration of the lysosomotropic drugs chloroquine or methylamine affect intestinal lipid metabolism. In the case of chloroquine, an excess of lipid as compared to controls was found in the mucosa during active lipid absorption without interfering with lipid transport into the lymph [33]. In the other case, a reduction in glycerol output by the intestine was found during lipid absorption [34]. In addition to this direct evidence, circumstantial evidence also points to a role for the lipase. The geographic localization of the enzyme to the proximal intestine in the villus tip cells is consistent with enzymes engaged in the absorption and transport of dietary constituents. Furthermore, in the life cycle of the rat, the enzyme is at its greatest activity during the neonatal period [35]. It is during suckling that the rat is exposed to the largest amount of dietary fat. Despite this information, a question of the quantitative importance of this enzyme to intestinal lipid metabolism is raised by the recent findings that although mucosal lipid is hydrolyzed at acidic pH, it is more quickly hydrolyzed at alkaline pH [36]. Furthermore, the amount of lipid in excess during the administration of chloroquine is less than would be expected if it were the primary enzyme responsible for the mucosal hydrolysis of dietary lipid. Therefore, the assessment of the physiological importance of intestinal acid lipase requires additional study. Acknowledgements This study was supported in part by National stitutes of Health Grant DK-38760 and in part Veterans Administration Medical Research Funds.
Inby
References Bernback, S., HemelI, 0. and Blackberg, L. (1985) Eur. J. Chn. Invest. 148, 233-238. Field, R.B. and Scow, R.O. (1983) J. Biol. Chem. 258,14563-14569. Tiruppathi, C. and Balasubramanian, A.S. (1982) Biochim. Biophys. Acta 712, 692-697. Moreau, H.. Gargomi, Y., Lecat, D., Junien, J. and Verger. R. (1988) B&him. Biophys. Acta 960, 286-293. Rao, R.H. and Mansbach, C.M., II (1989) Biochim. Biophys. Acta 1043, 273-281. Warner, T.G., Dambach, L.M., Shin, J.H. and O’Brien, J.S. (1981) J. Biol. Chem. 256, 2952-2957. Sjoberg. E.R., Hatton, J.D. and O’Brien, J.S. (1987) B&hem. J. 248, 139-144.
26 8 Burrier, R.E. and Brecher, P. (1983) J. Biol. Chem. 258, 1204312050. 9 Brown, W.J. and Sgoutas, D.S. (1980) B&him. Biophys. Acta 617, 305-317. 10 Teng, M. and Kaplan, A. (1974) J. Biol. Chem. 249, 1064-1070. 11 Imanaka, T., Muto, K., Ohkuma, S. and Takano, T. (1984) J. B&hem. 96. 1089-1101. 12 DeWolf, M., Lagrou, A., Hilderson, H.J. and Dierick, W. (1978) Lipids 13, 598-604. 13 Burton, B.K. and Mueller, H.W., Jr. (1980) B&him. Biophys. Acta 618, 449-460. 14 Sando, G.N. and Rosenbaum, L.M. (1985) J. Biol. Chem. 260, 15186-15193. 15 Hanosh, M., Ganot, D. and Hamosh, P. (1979) J. Biol. Chem. 254, 12121-12125. 16 Roberts, I.M. and Jaffe, R. (1986) Gastroenterology 90,1170-1175. 17 Knauer, T.E. and Weglicki, W.B. (1983) Biochim. Biophys. Acta 753, 173-185. 18 Rindler-Ludwig, R., Patsch, W., SaIier, S. and Braunsteiner, H. (1977) B&him. Biophys. Acta 488, 294-304. 19 Laemmh, U.K. (1970) Nature 227,680-685. 20 Folch, J., Lees, M. and Sloane-Stanley, G.H. (1957) J. Biol. Chem. 266.497-509. 21 Skipski, V.P., Peterson, R.F. and Barclay, M. (1964) Biochem. J. 90, 374-378. 22 Lowry, O.H., Rosebrough, N.J., Farr, A.L. and Randall. R.J. (1951) J. Biol. Chem. 193, 265-275. 23 Moreau, H.. Gargouri, Y., Lecat, D., Junien, J.L. and Verger, R. (1988) B&him. Biophys. Acta 959, 247-252.
24 Momsen, 384-388.
W.E. and
Brockman,
H.L.
(1976) J. Biol. Chem.
251.
25 Borgstrom, B. (1954) Biochim. Biophys. Acta 13, 149-150. 26 Vahouny, G.V., Weersing, S. and Treadwell, C.R. (1965) Biochim. Biophys Acta 98, 607-616. 27 Haley, N.J., Fowler, S. and De Duve, C. (1980) J. Lipid. Res. 21, 961-969. 28 Jensen, G., Daggy, B. and Bensadoun. A. (1982) B&him. Biophys. Acta 710, 464-470. 29 Mattson, F.H. and Beck, L.W. (1956) J. Biol. Chem. 219, 735-740. 30 Entressangles, B., Sarl, H. and Desnuelle, P. (1966) Biochim. Biophys. Acta 125, 597-600. 31 Mooney, R.A. and Lane, D.M. (1981) J. Biol. Chem. 256, 1172411733. 32 Partin, J.C., Mercer, T.R. and Schubert, W.K. (1986) in Proceedings of the 26th Annual Meeting of the Electron Microscopic Society (Archenaux, C.J., ed.), pp. 194-195, Baton Rouge, LA, Claitor’s. 33 Mansbach, C.M., Arnold, A. and Garrett, M. (1987) Am. J. Physiol. 253, G673-G678. 34 Hulsman, W.C., Stan, H. and Breeman, W.A.P. (1981) Biochem. Biophys. Res. Commun. 102, 440-448. 35 Coats, P.M., Brown, S.A.. Jumanvan, J. and Koldovsky, 0. (1977) B&hem. J. 331-338. 36 Tipton, A.D., Frase, S. and Mansbach, C.M., II (1989) Am. J. Physiol. 257. G871-878. 37 Laurent, T.C. and Killander, J. (1964) J. Chromatogr. 14, 317-330.
BBALIP
19
Acta, 1046 (1990) 19-26
53458
Purification and partial characterization of intestinal acid lipase R. Hanumantha
Rao and Charles M. Mansbach,
II
Department of Medicine, Division of Gastroenterology, The University of Tennessee, Memphis and The Veterans Administration Medical Center, Memphis, TN (U.S.A.)
(Revised
Key words:
Lipase;
Intestine;
(Received 23 January 1989) manuscript received 18 April 1990)
Lipolysis;
Protein purification;
Intestinal
mucosa;
Lipid metabolism
Intestinal acid lipase is an enzyme whose greatest specific activity is localized to the villus tips of the proximal intestine (Rae, R.H. and Mansbach, C.M. (1990) Biochim. Biophys. Acta 1043,273-280). This suggests that it plays a role in the processing of dietary lipids. We purified the enzyme in order to better characterize it. Acid lipase was isolated from intestinal mucosa of rats by a combination of ammonium sulfate precipitation, butanol extraction and chromatography on DEAE Bio-Gel, CM Bio-Gel and Sephadex G-75. This resulted in a single protein of M, 53700 on SDS-polyacrylamide gel electrophoresis. The isolation scheme produced a 3344-fold purification resulting in an enzyme whose specific activity was 801 pmol/min per mg protein. The yield was 50%. The purified enzyme was stimulated @&fold) by the addition of tauro- or glycocholate but no other conjugated bile acid. A sharp peak in activity occurred at pH 5.6. The pl of the enzyme was 6.2. The reaction products produced under prolonged incubation suggested that monoacylglycerol was not hydrolyzed since an overabundance of monoacylglycerol was found with respect to the amount of fatty acid produced. These results suggested that intestinal acid lipase is potentially important in the metabolism of dietary lipids. Its proportionate role awaits further documentation.
Introduction A number of acid active Iipases, usually lysosomal in origin, have been purified from various tissues and species. The activity of these Iipases varies from pmol to nmol/mg protein per min. Those lipases with the greatest activity are secreted into the gastrointestinal tract. These include the pharyngeal [l], lingual [2] and gastric lipases [3,4]. Those with the least activity are the ones isolated from tissues such as the fibroblast [4] which normally do not process much lipid. We have recently studied a similar lipase of intestinal origin [5]. This hpase appeared to be in position to play a role in the intestinal metabolism of absorbed lipids based on the circumstantial evidence that it is found in greatest activity in the proximal intestine in the villus tips. This is the location expected for enzymes and transporters engaged in the absorption and processing of dietary substrates. The quantitative role of this enzyme in lipid metabolism remains to be explored, however.
Correspondence: see, Memphis, 38163, U.S.A.
Charles M. Mansbach, II, The University of Tennes951 Court Avenue, Room 555 Dobbs, Memphis, TN
0005-2760/90/$03.50
0 1990 Elsevier Science Publishers
B.V. (Biomedical
The current investigations were conducted toward purifying acid lipase from the intestine. Previously purified acid Iipases [l-4,6-18] have shown remarkable similarity with regard to their pH optima, their molecular weight and their pl, but not their activity against long acyl chain (C > 12) triacylglycerol (TG) substrate. We wished to determine if the intestinal enzyme were similar to those previously reported with respect to these physicochemical characteristics. Equally important, with respect to considerations for a physiological role of the enzyme, is the specific activity of the purified enzyme. If high, it would more likely play a role in intestinal lipid metabolism. Methods Tissue preparation Male Sprague-Dawley rats weighing 250-350 g were used. Usually 16 rats were processed at a time. Under ether anesthesia, their intestines were removed to an iced glass plate. The mucosa from the proximal 3/4 intestine was harvested by the aid of giass microscope slides and placed in Buffer A (taurocholate, 4 mM; dithiothreitol (DTT), 2.5 mM; Tris-HCl, 10 mM (pH 7.4). The mucosa was homogenized in 10 wt/v Buffer A using a gIass homogenizer with Teflon pestle. The hoDivision)
20 mogenate was centrifuged for 3. lo5 g. min in a Sorval RC 5C centrifuge (DuPont Instruments, Norwalk, CT). The pellet was extracted three times with 50 ml of Buffer A and re-centrifuged in between. The supernatants were combined and adjusted to 50% saturated (NH4)*S04 and stirred gently at 2°C for 30 min. The mixture was then centrifuged for 3.10’ g. min. The resulting pellet was resuspended in Buffer A supplemented with 20% ethylene glycol (Buffer B) and dialyzed overnight with two changes of buffer. The dialysate was extracted with butanol at 2°C which was added slowly until it reached 0.5 volumes. The mixture was stirred for 20 min and then centrifuged for 8. lo4 g. min. The butanol was removed by suction and the lower, aqueous layer re-extracted with butanol as previously. The proteinaceous material that forms at the butanol/H,O interface was discarded. The combined aqueous phases was dialyzed against two changes of buffer B. Chromatographic procedures The dialysate was applied to a DEAE Bio-Gel A (100-200 mesh, Bio-Rad, Richmond, CA) column (1.5 x 24 cm) without prior concentration. The column was pre-equilibrated with Buffer B and eluted with the same buffer. Active fractions were concentrated on a YM 10 membrane (Amicon Corp, Danvers, MA) and dialyzed against Buffer C (sodium acetate, 20 mM; dithiothreitol (DTT), 2.5 mM; ethylene glycol, 20% and taurocholate, 4 mM; pH 5.5). The buffer was changed three times. The dialysate was placed on a CM Bio-Gel (Bio-Rad Laboratories, 50-200 mesh) column of 1.5 x 24 cm. The column was pre-equilibrated with Buffer C and eluted with a NaCl gradient of 0 to 0.4 M using a gradient mixer. The flow rate was 4.8 ml/h. 2-ml fractions were collected. The active fractions were dialyzed against Buffer B with three changes of dialysand and concentrated on a YM 10 membrane (Amicon Corp). The concentrated material was applied to a Sephadex G-75 superfine column (Pharmacia Fine Chemicals, Piscataway, NJ) of 2.5 X 65 cm and eluted with Buffer B. The column flow rate was 6 ml/h. 2-ml fractions were collected. The column was calibrated using bovine serum albumin, 66 kDa; ovalbumin, 45 kDa; chymotrypsinogen, 25 kDa; and ribonuclease A, 13.7 kDa (all supplied by Sigma Chemical, St. Louis, MO). All chromatographic procedures were performed in a cold room. SDS polyacrylamide gel electrophoresis (SDS PAGE) was performed in a Hoefer model 500 apparatus (Hoefer Scientific Instruments, San Francisco, CA) as suggested by Laemmli [19]. Standard proteins of known M, (see legend to Fig. 3) or samples of acid lipase were heated to 100°C for 4 min in the presence of SDS, 2% and /3-mercaptoethanol, 5%. 20 pg of acid lipase was applied to the 10% cross-linked gel and electrophoresed for 5 h using approx. 30 mA per gel. After development, the gel
was stained with 0.1% Coomassie Blue R-250 in 40% methanol and 10% acetic acid. It was destained with 10% methanol and 7.5% acetic acid. Isoelectric focusing was performed using a LKB 110 ml column (LKB Washington, DC) cooled to 2” C. The ampholyte used was Pharmalyte (Pharmacia, Uppsala, Sweden) which was used at a final concentration of 2% at pH 4.0-6.5. A gradient of O-40% glycerol was constructed and the enzyme applied to the middle of the column in 0% glycerol. The protein used was the dialyzed soluble extract (see Table I). The total run was 18 h at 1600 V. Protein modifying reagents The purified enzyme was preincubated for 20 min in the presence of inhibitor at 37” C. Acid lipase activity remaining was assayed for 2 min as in Analytical Methods. The following potential inhibitors were utilized (Sigma Chemical): N-ethylmaleimide, N-bromosuccinimide, 2,4,6-trinitrobenzenesulfonic acid, diethyl pyrocarbonate and diethyl p-nitrophenyl phosphate (E600). Substrate specificity studies Purified enzyme was incubated in the presence of either glyceryl tri[i4C]oleate, l-palmitoyl-2-[‘4C]oleoylphosphatidylcholine (New England Nuclear), cholesteryl [i4C]oleate (New England Nuclear), 4-methylumbelliferyl oleate or, 4-methylumbelliferyl palmitate (both from Sigma Chemical) in the taurocholate containing buffer described in Analytical Methods. The incubations were for 2 min each at 3 mM substrate. Released oleate from TO and cholesteryl oleate were quantified as in Analytical Methods. Potential phospholipase A, or A, activity was determined by extracting the incubation mixture [20] and separating the products on TLC using as the mobile phase chloroform/ methanol/ acetic acid/H,0 (25 : 15 : 4.2, v/v) [21]. Lyso derivatives and fatty acids, the potential products, were identified by authentic standards. The appropriate areas were scraped from the plate and radioactivity determined (Analytical Methods). The production of umbelliferone was quantified by fluorometry [6] with a Hitachi model 650-15 fluorecence spectrophotometer (Hitachi, Tokyo, Japan) using 320 nm as the excitation wavelength and 450 nm as the emission wavelength. Analytical methods Acid lipase was assayed using glyceryl tri[14C]oleate (TO) (New England Nuclear, Boston, MA), 3 mM (1 . lo5 dpm/pmol); sodium taurocholate, 20 mM (Calbiochem, La Jolla, CA); DTT, 1 mM; glycerol, 0.8 M; sodium acetate, 50 mM (pH 5.8) and appropriate amounts of protein depending on the degree of purity in a total volume of 250 ~1 [5]. The TO was prepared for assay by sonifying it in 4% glycerol and 4 mM tauro-
21 cholate. The reaction was stopped and released oleate extracted by the addition of 0.5 ml carbon tetrachloride/ hexane (2 : 1, v/v) containing 1 mM Triton X-100 and 0.1 mM oleate, and 0.75 ml of ethanol/ water (3 : 1, v/v) containing 70 mM NaOH [6]. Radioactivity was determined using Scintiverse II (Fisher Scientific) in a Packard 1500 liquid spectrometer (Packard Instruments, Downers Grove, IL). 96% of the extracted radioactivity was FFA. The Folch [20] extraction procedure was tested for its ability to extract the total FFA present by extracting TO (750 nmol) plus a known amount of [‘4C]oleate (250 nmol). 98.8% of the dpm were in the organic phase. 85% of lipid dpm after a lipase incubation that were subjected to TLC were recovered from the plate as compared to the direct determination of radioactivity. Protein was determined by the method of Lowry et al. [22]. Thin-layer chromatography (TLC) was performed on Silica-gel H layers using the solvent system hexane/ diethyl ether/ methanol/ acetic acid (80 : 20 : 6 : 2, v/v).
Results The steps used in the purification of intestinal acid lipase are shown in Table 1. It is evident that solubilization of the enzyme into taurocholate micelles enhanced its activity as compared to that present in the whole homogenate. This resulted in more enzymatic activity than was originally present in the non-processed homogenate. This may have been due to the solubilized enzyme having a preferred conformation for interaction with substrate. Alternatively, an inhibitor could have been present which pelleted on centrifugation. Multiple
other detergents were tried in attempts to solubilize the enzyme as previously reported [5]. All of these inhibited enzyme activity [5]. A O-50% ammonium sulfate precipitation step was used to concentrate the enzyme and to remove soluble proteins. This resulted in a 3.8-fold purification (Table I) which is at the upper end of the expected range (2-4-fold). The resulting material had moderate turbidity and could not be loaded onto chromatographic columns. In order to more completely solubilize the enzyme and to remove associated membranous fragments, a butanol extraction step was employed. 0.5 vol. of butanol (2” C) was added slowly while stirring which was continued for 20 min. The mixture was centrifuged for 38 000 x g. min to separate the phases and the butanol layer removed by suction and discarded. The cake which formed at the butanol/aqueous interface contained 24% of the enzymatic activity. The interfacial proteins were extracted with Buffer A. Only 10% of the activity could be recovered. Therefore, the interfacial proteins were discarded as well. The aqueous phase was extracted a second time with butanol and the phases separated as before. The aqueous phases were combined. The combined aqueous phases had an increase in specific activity of only 1.6-fold (Table I) as a result of the butanol treatment. This step was necessary, however, to yield a clear solution which could be chromatographed. The aqueous phase from the butanol extract was applied to a DEAE Bio-Gel column (1.5 X 24 cm) after dialysis against buffer A. It was eluted with the same buffer. The enzyme did not bind to the column. Nevertheless, considerable unwanted proteins did which resulted in a 6.4-fold (Table I) purification. Since no binding occurred, the data are not presented further.
TABLE I Purification of intestinal acid lipase Fraction
Total protein
(mg)
Specific activity ( pmol OA/min per mg protein”)
Total activity ( pmol OA/min)
Recovery b (W)
Purification (fold)
Whole homogenate
2820
0.240
616
100
1
Soluble extract
1585
0.611
968
143
2.5
155
9.4
Ammonium
sulfate PPtn’
Butanol extraction DEAE Bio-Gel
463
2.26
1047
234
3.36
785
116
14.0
827
122
88
39.3
21.0
CM B&Gel
2.30
206
474
70
861
Sephadex G-75
0.418
801
335
50
3344
a Oleic acid rekased. b Percentage recovery is based on that present in the whole homogenate. ’ Precipitation.
22 CHROMATOGRAPHY
OF ACID
CHROMATOGRAPHY
LIPASE ON CM SIOGEL
LIPASE
12
= _E
3
mm*-
44
I_
OF ACID
ON SEPHADEX
G-75
PROTEIN
54
64
?b
0
FRACTION FRACTION
NUMBER
Fig. 1. The proteins which passed through the DEAE Bio-Gel column without adhering to the support were chromatographed on CM BioGel. Acid lipase activity (solid tine) was eluted from the column in response to an NaCl gradient as indicated by the open chain line. Proteins, measured at A,,, are indicated by the dashed line.
The active fractions were concentrated on a YM 10 membrane and dialyzed against buffer B. The enzyme was then fractionated on CM Bio-Gel (1.5 x 24 cm) previously equilibrated with the same buffer. 2-ml fractions were collected. After 30 ml of buffer had passed through the column, a NaCl gradient elution was established with a maximal concentration of 0.4 M. As shown in Fig. 1, at 0.05 M NaCl the enzyme began to elute from the column. Fractions 61-74 were pooled, dialyzed against buffer A and concentrated on a YM 10 membrane. A 9.8-fold purification was obtained (Table I>* The concentrated sample was next separated on Sephadex G-75 superfine using a column 2.5 X 65 cm. The enzyme was eluted with buffer A as a single peak coincident with the protein peak as shown in Fig. 2. Calibration of the column with proteins of known molecular weight suggested a hydrodynamic radius consistent with a M, of 44000. Fractions 67-77 were pooled and concentrated on a YM 10 membrane. A further 4-fold purification was achieved (Table I). In sum, the six purification steps gave a 3344fold purification with a 50% yield based on activity present in the original homogenate. SDS-PAGE was performed on the concentrated G-75 eluent. The results are shown in Fig. 3. By Coomassie blue stain, only one band was visible. No other bands were visualized with silver staining (data not shown). When compared to other proteins of known molecular weight which were similarly electrophoresed, acid lipase had a suggested M, of 53700. It is not clear why the
NUMBER
Fig. 2. Fractions 61-74 from the CM Bio-Gel column were concentrated and passed over a Sephadex G-75 column. Acid lipase activity (solid line) eluted as a single peak with a trailing shoulder. Proteins, measured at A,, and shown by the dashed line, peaked in a coincident manner with the major protein peak. The column was calibrated by chromatographing proteins of known molecular weight. These were: bovine serum albumin (M, 66000), ovalbumin (M, 45000). chymotrypsinogen (M, 25000) and ribonuclease A (M, 13 700). The molecular weight of acid lipase was calculated by plotting the K,, [37] of the known proteins against their molecular weights.
66 KD 45 KD 36 KD 29 KD 24 KD
Fig. 3. Fractions 66-77 from the Sephadex column were concentrated and subjected to polyacrylamide gel electrophoresis in the presence of SDS. The protein migrated as a single band of M, 53700. This was calculated by plotting the R, values of the known proteins against their molecular weights as a semilog plot. These proteins were: bovine serum albumin (M, 66000), ovalbumin (M, 45000), glyceraldehyde3-phosphate dehydrogenase (M, 36000), carbonic anhydrase (M, 29000) and trypsinogen (M, 24000).
23
ISOELECTRIC
FOCUSING
OF ACID
LIPASE
TABLE
II
Effect of various bile salts on intestinal acid lipase activity =
14-
E
.
-.-.‘D--
OD260
+
ACTIVITY
Activity OA b released (nmol/min) ’
Bile salt a
No addition
2.6f0.3
Taurocholate
48.7 f 4.1 3.5 *0.4
Glycodeoxycholate
50.7 + 4.7
Glycocholate
4
2
6
10
6
PH Fig. 4. Isoelectric focusing of acid lipase is shown in the figure. Acid lipase activity is shown by the solid line and protein, as measured at GD,se, by the dashed line. Soluble extract (Table I) was used as the protein source. A pH gradient of 4.0-6.5 was used in the presence of a glycerol gradient (O-401).
enzyme appears to have a molecular weight which is smaller on gel permeation chromatography than when it is chromatographed as the denatured, reduced protein. This has been observed previously with calf pharyngeal lipase [l]. One possibility is an interaction of the enzyme with the gel. This could lead to a spuriously
EFFECT
OF pH ON ACID
LIPASE
ACTIVITY
600-
Glycochenodeoxycholate
6.5 f 0.9
Taurodeoxycholate
6.9kO.8
Taurochenodeoxycholate
6.1 f 0.8
a All bile acids were present at a concentration of 20 mM. b Oleic acid released. ’ The data are the meanfS.E. of four experiments. In each case, blank tubes containing the appropriate bile acid were run in parallel with the tubes containing enzyme.
delayed appearance of the enzyme in the chromatogram which would yield a result suggesting a smaller protein than what was actually present. Isoelectric focusing of the enzyme was performed and the results shown in Fig. 4. A p1 of 6.2 is suggested. The activity yield from this procedure was low so that it was not useful as part of the purification scheme. Acid lipase activity as a function of pH is reported in Fig. 5. A sharp rise in activity is seen between pH 5.2 and 5.6 which is the pH at which maximum activity of the purified enzyme occurs. Enzymatic activity progressively fell at more alkaline pH but persisted into the alkaline range. The activity at pH 8 was, however, 7.5fold less than at pH 5.6.
WO-
TABLE
III
400-
Identification
of reaction products of intestinal acid lipase
soo-
TG
DG
MG
2uo-
Control (no enzyme)
57630
1033
100 -
Experimental a (+ enzyme)
14983
11756
Net change 4.0
5.0
6.0
7.0
6.0
PH Fig. 5. Acid enzyme was 4.0-5.6 and buffers were 5.8) normally
lipase activity as a function of pH is shown. Purified used. Acetate buffer, 0.05 M, was used between pH Tris-maleate buffer, 0.05 M, between pH 5.6-8.4. These used as a substitute for the sodium acetate, 0.05 M (pH present in the assay. The other components of the assay remained the same (see Methods).
Ratio
- 42 647
FFA
477
13167
+10723
18179
+12690 1
2 322
+ 15 857 1.2
1.5
a 0.51 pg pure enzyme was incubated with 750 nmol [‘?I TO (spec. act. 99.8 dpm/nmol, 20 mM taurocholate and 50 mM sodium acetate buffer (pH 5.8) in a total volume of 0.25 ml. The incubation was for 1 h at 37OC in a shaking water bath. The reaction was stopped by the addition of chloroform/methanol (2: 1, v/v) and the lipids extracted [20]. Control incubations were the same except no enzyme was added. The data are the mean of two separate experiments performed in duplicate. Values are in dpm.
24 TABLE
IV
Substrate
a
Glycerol
Fatty acid released b (nmol/min per mg protein)
[i4C]oleate
692
l-Palmitoyl-2-[14C]oleoylphosphatidylcholine
0
Cholesteryl
0
[‘4C]oleate
4-Methylumbelliferyl
oleate
4-Methylumbelliferyl
palmitate
12519 1824
a Each substrate was present in a concentration of 3 mM. b Acid lipase activity was determined as in Materials and Methods. All assays were performed twice in duplicate.
Of particular interest with respect to a mucosal enzyme is the effect of bile salts on enzymatic activity. The purified enzyme was significantly stimulated by trihydroxylated bile salts as shown in Table II. Both glyco- and taurocholyl conjugates were equally effective in increasing enzyme activity. These bile acids stimulated enzymatic activity 20-fold. These data differ significantly with that previously reported in impure preparations which are stimulated by a variety of bile acids
PI. In Table III are shown data with regard to the reaction products produced from TG upon prolonged lipolysis. As shown in the table, 74% of the TG originally present was hydrolyzed after 1 h incubation at 37 o C. As was observed with the crude preparation, an excess of MG was observed for the amount of FFA released. For each DG, one FFA and for MG two FFA should be produced. It is immediately apparent that there is less FFA than would be expected for the amount of DG and MG present. Although the reason
TABLE
V
Effect of protein-modifying
reagents on acid lipase activity
Acid lipase was preincubated for at 37OC. Glyceryl trioleate was for 2 min. Acid lipase activity Materials and Methods. Control was 692 pmol acid released/mm performed twice in duplicate.
20 min in the presence of the reagent added and the incubation continued was determined as described under activity with no addition of reagent per mg protein. All assays were
Reagent
mM Reagent
Activity
No addition
_
100
N-Ethylmaleimide
4.0
6
N-Bromosuccinimide
4.0
113
4.0
49
4.0
100
0.1
36
2,4,6-Trinitrobenzenesulfonic Diethyl
pyrocarbonate
Diethyl
p-nitrophenyl
a Percentage
of control
acid
phosphate values.
(E600)
a
for this is not clear, it is evident that the reaction is likely to stop at MG and that complete hydrolysis of TG does not occur. The substrate specificity of the purified lipase was investigated by its incubation with a variety of potential substrates. The results are shown in Table IV which reveal that it is inactive against cholesteryl oleate and phosphatidylcholine. It is, however, even more active against the umbelliferone esters than against TO. This has been noted previously with other lipases [6,13,14]. The results of inhibitor studies are shown in Table V. These demonstrate that there is likely to be a serine at the active site of the enzyme as both N-ethylmaleimide and E-600 were greatly inhibitory. Partial inhibition was observed using large concentrations of trinitrobenzenesulfonic acid indicating that a lysine residue is near but not at the active site. Inhibitors of histidine (diethyl pyrocarbonate) and tryptophane (bromosuccinimide) did not disturb enzymatic activity indicating that these two amino acids are not near L active site. Discussion We have purified a mucosal, acid active, lipase of M, 54 kDa. This lipase is very active, second only in activity to the acid lipase isolated from the rabbit stomach [4] when TO is used as substrate. It is lysosomal in origin and its maximal activity is in the villus tip cells of the proximal intestine [5]. It is distinguished from preduodenal lipase (lingual lipase in the rat [23]) by its continued activity despite esophageal diversion for 48 h [5] and from pancreatic lipase both by its stimulation by taurocholate rather than its inhibition [24] and by its pH optimum [25]. The mucosal acid lipase isolated is remarkably similar to a variety of other acidic lipases isolated from multiple species and tissues [l-4,5-18]. Of the 17 different reports, the average (*SD.) M, is 53300 _t 1100 excluding the human placental enzyme [13]. The pH optimum of the lipases average 4.8 + 0.5. This average is slightly more acidic than the 5.6 of the mucosal enzyme reported here. Nevertheless, there are five lipases whose pH optima either overlap with the intestinal enzyme or are within 0.3 pH units of it [2-4,12,15,18]. Intestinal acid lipase is acidic as suggested by its pl of 6.2. This is comparable to the p1 of the hepatic enzyme [9] and slightly more acidic than the cardiac myosidic and calf pharyngeal enzymes whose pl are 6.3 and 7.0, respectively (17.1). The similarity of the acid lipases suggests that considerable sequence homology will be identified. Despite these apparent similarities, differences between the intestinal enzyme and other acidic lipases are evident. In response to bile acids, the rat lingual enzyme has been shown to have both increased [2] and decreased [16] activity; the human gastric lipase is inhibited by all bile acids [3]. By contrast, the intestinal
25 enzyme is significantly stimulated by trihydroxylated bile acids. In this regard it is similar to sterol ester hydrolase which is specifically stimulated by taurocholate [26] and acid cholesterol esterase which is also stimulated by taurocholate [27]. With regard to the non-ionic detergent, Triton X-100, the liver enzyme is consistently stable [1,7-91 whereas the intestinal enzyme is completely inhibited. The co-addition of taurocholate partially restores activity of the intestinal enzyme in response to Triton treatment, however [5]. In sum, these data would suggest that the intestinal lipase has an interfacial recognition site which is similar to the pancreatic and cholesterol esterases but different from the acid lipases. The active site of the intestinal enzyme is likely to differ from that of the sterol ester hydrolases in that the lipase does not hydrolyze cholesterol esters. Its inability to hydrolyze phosphatidylcholine also distinguishes it from hepatic lipase which does [28]. The degree to which acid lipases hydrolyze a TG substrate also varies. The rat lingual enzyme completely hydrolyzes TG [2,15] whereas the liver enzyme is not able to hydrolyze monoacylglycerol [6,8]. In this regard, the liver and intestinal acid lipases are similar to pancreatic lipase whose inability to completely hydrolyze TG, leaving monoacylglycerol, is well established [29,30]. The purified intestinal acid lipase is a very active enzyme, with a specific activity of 801 pmol/min per mg prot. This is the most active of all reported acidic lipases with the exception of the rabbit gastric lipase [4]. In general the acid lipases can be divided into three groups according to their specific activities. The lipases with the most activity are those which are secreted into the intestinal lumen. In this group are the gastric juice lipases from both human [3] and rabbit [4]. Both of these have activities similar to the intestinal enzyme. The other lipases in this group are the lingual lipases which also demonstrate considerable activity [1,2] with the exception of one isolation [15]. Although the intestinal enzyme is as active as the gastric and lingual ones there is no evidence that it is secreted into the lumen. The intestinal enzyme has a subcellular localization in lysosomes [5]. As such it would make its extracellular secretory potential unlikely. In the second group with intermediate activity is the liver enzyme. Four different isolates demonstrate activity in the range of 2.5 to 7.1 ~mol/min per mg prot. [6-8,111. One isolate, however, is much more active (408 ~mol/min per mg prot.) [9]. The liver enzyme appears to be involved in the hydrolysis of TG stored in the hepatocyte [31]. Finally, there is a third group of lipases whose activity is quite low, in the nanomolar range. These lipases have been isolated from organs that normally process little lipid such as the thyroid [12], fibroblasts [14], placenta [13], cardiac myocytes [17] and leukocytes [18].
The high specific activity of intestinal acid lipase would suggest that it does in fact play a role in the normal intestinal processing of dietary lipid. This potential is enhanced by the finding that in neonates who congenitally lack acid lipase activity in their intestines large lipid droplets are found at electron microscopy [32]. Additional direct evidence comes from experimental studies in rats in which it has been shown that the administration of the lysosomotropic drugs chloroquine or methylamine affect intestinal lipid metabolism. In the case of chloroquine, an excess of lipid as compared to controls was found in the mucosa during active lipid absorption without interfering with lipid transport into the lymph [33]. In the other case, a reduction in glycerol output by the intestine was found during lipid absorption [34]. In addition to this direct evidence, circumstantial evidence also points to a role for the lipase. The geographic localization of the enzyme to the proximal intestine in the villus tip cells is consistent with enzymes engaged in the absorption and transport of dietary constituents. Furthermore, in the life cycle of the rat, the enzyme is at its greatest activity during the neonatal period [35]. It is during suckling that the rat is exposed to the largest amount of dietary fat. Despite this information, a question of the quantitative importance of this enzyme to intestinal lipid metabolism is raised by the recent findings that although mucosal lipid is hydrolyzed at acidic pH, it is more quickly hydrolyzed at alkaline pH [36]. Furthermore, the amount of lipid in excess during the administration of chloroquine is less than would be expected if it were the primary enzyme responsible for the mucosal hydrolysis of dietary lipid. Therefore, the assessment of the physiological importance of intestinal acid lipase requires additional study. Acknowledgements This study was supported in part by National stitutes of Health Grant DK-38760 and in part Veterans Administration Medical Research Funds.
Inby
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