Enhanced extracellular enzymatic peptide hydrolysis in the sea-surface microlayer

Marine Chemistry 73 Ž2001. 319–332 www.elsevier.nlrlocatermarchem

Enhanced extracellular enzymatic peptide hydrolysis in the sea-surface microlayer Marina Kuznetsova) , Cindy Lee Marine Sciences Research Center, State UniÕersity of New York, Stony Brook, NY 11794-5000, USA Received 19 January 2000; received in revised form 17 November 2000; accepted 27 November 2000

Abstract The accumulation of dissolved organic matter ŽDOM. at the air–sea interface is controlled by dynamic physical processes at the boundary between ocean and atmosphere. Much of the DOM concentrated in the surface microlayer is thought to be protein or glycoprotein. Enzymatic hydrolysis of these and other biopolymers is an important step in the microbial uptake of dissolved and particulate organic matter in many aquatic environments. We employed a sensitive fluorescence technique to investigate differences between extracellular enzymatic peptide hydrolysis in the sea surface microlayer and corresponding subsurface water from Stony Brook Harbor, NY. We separated the microlayer from its underlying water and thus measured hydrolysis potential rather than an in-situ process. Peptide turnover was always faster in the microlayer than in subsurface waters. This was confirmed by allowing a new surface film to form on subsurface water; hydrolysis was still faster in the new surface film. In a year-long study, we found the relative difference between turnover times in the surface film and subsurface waters to vary greatly with season. While rate constants of peptide hydrolysis were generally higher in both microlayer and bulk water samples in springrsummer than in fallrwinter, the difference in activity between the two environments was greatest in winter. Enhanced hydrolysis in the sea surface microlayer is likely due to the greater concentrations of DOM in the microlayer. Seasonal changes in distribution of hydrolytic activity between surface film and subsurface water probably reflect seasonal variation in the mechanisms of DOM enrichment, which depend on water temperature, substance and energy fluxes across the water–air boundary, activity of aquatic organisms and other seasonal variables. q 2001 Elsevier Science B.V. All rights reserved. Keywords: Peptide hydrolysis; Sea-surface microlayer; Microlayer enrichment; THAA

1. Introduction The thin layer of water at the air–sea interface represents a unique physico-chemical habitat for ma-

)

Corresponding author. Tel.: q1-516-632-8741; fax: q1-516632-8820. E-mail addresses: [email protected] ŽM. Kuznetsova., [email protected] ŽC. Lee..

rine biota. The microlayer is subject to constant alteration by a number of dynamic and non-equilibrium processes that take place at the air–water interface. However, there are consistent characteristics that define the special nature of the layer. Both dissolved and particulate materials accumulate in the microlayer with enrichment factors up to 10 3 ŽLiss and Duce, 1997.. A remarkable feature of the surface layer is that different constituents concentrate in the microlayer non-uniformly; consequently, the ratios

0304-4203r01r$ - see front matter q 2001 Elsevier Science B.V. All rights reserved. PII: S 0 3 0 4 - 4 2 0 3 Ž 0 0 . 0 0 1 1 6 - X

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of many dissolved elements differ here from those in bulk water ŽHorne, 1969.. The distribution of organic matter between dissolved, colloidal and particulate phases in the microlayer water is also different from that in bulk water. The sea surface microlayer is the only area of the water column where significant Žup to 10008rm. temperature gradients may be present ŽKhundzua et al., 1977.. Thus, although the sea surface is an open system, it represents a distinctive and complex marine environment. Enrichment of proteinaceous material and other constituents in the surface microlayer is well documented ŽLiss and Duce, 1997.. Natural organic components appear to be the principal film-forming components in the surface microlayer, with glycoprotein-dominated material as the major organic component, although lipid compounds can dominate in samples collected from polluted waters ŽBaier et al., 1974; Williams et al., 1986.. Rates of biochemical transformations are often enhanced in the sea surface microlayer relative to underlying water ŽHenrichs and Williams, 1985; Williams et al., 1986; Carlucci et al., 1992., although few significant correlations have been observed between these rates and chemical and biological parameters ŽWilliams et al., 1986.. Hydrolysis of biopolymers is thought to be a rate-limiting step in microbial uptake of dissolved and particulate organic matter in many aquatic environments ŽHoppe, 1986; Halemejko and Chrost, ´ 1986; Meyer-Reil and Koster, 1992.. Recently a ¨ novel approach to studying extracellular peptide hydrolysis rates in natural environments was developed by Pantoja et al. Ž1997. and applied to seawater and sediments in a coastal salt marsh ŽPantoja and Lee, 1999.. This technique employs a fluorescently labeled peptide as a substrate for peptide hydrolysis. The suitability of these synthetic peptides as analogs of natural compounds has been evaluated in terms of their adsorptive properties, fluorescent attributes, and the absence of their incorporation across cell membranes ŽPantoja et al., 1997.. Here we apply this method to analyze biological, physical, and chemical factors controlling differences in the capacity for enzymatic peptide hydrolysis between the sea surface microlayer and the corresponding subsurface water. We compare laboratory-measured hydrolysis rate constants in the microlayer with those in the

water below over 1 year, and relate seasonal changes in the distribution of hydrolytic activity to dynamic processes occurring at the air–water interface.

2. Materials and methods 2.1. Sampling of the natural microlayer and subsurface water Water samples were collected from Main Channel in Stony Brook Harbor, NY, around 9 a.m. Žexcept on 12r7r1998 when collections were at 2 p.m.. between October 1997 and February 1999 ŽTables 1 and 2.. Sampling was conducted on calm days during high tide upcurrent from a small platform that stuck out about 2 m into the channel. Although it was difficult to determine the extent of surface slick at this location, the site provided easy access to relatively clean water. Microlayer samples ŽML. were taken with a polyester-mesh screen Ž1-mm mesh openings. attached to a high-density polyethylene frame 30.5 mm in diameter. The screen was immersed vertically, rotated 908 and the surface film lifted off ŽGarrett, 1965.. The thickness of the microlayer sample collected using this technique is about 200–400 mm. The microlayer was collected by shaking the screen over a straight-sided polypropylene jar with an opening smaller than the screen frame to prevent water adhering to the frame from contaminating the sample. Subsurface Žor bulk. water samples ŽSS. were collected by submerging a polypropylene bottle by hand 15 cm Ž"2 cm. below the sea surface; the bottle was opened and closed in place to avoid surface microlayer interference. Subsamples of ML and SS water were immediately placed in glass bottles and preserved with formaldehyde for bacteria counts. Subsamples for measurement of dissolved total hydrolyzable amino acids ŽTHAA. concentrations were filtered Ž0.2 mm Nuclepore. and frozen until analysis. 2.2. Measurement of peptide hydrolysis Natural microlayers are constantly renewing and mixing with underlying water, so that microlayer composition at any sampling time is the result of a

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321

Table 1 Effect of sample storage on extracellular LYA-tetraalanine hydrolysis turnover time. Hydrolysis was measured at 22.58C Storage temperature and time

Sampling date and water temperature

Source

t fresh

Žt SS rt ML . fresh

tstored

Žt SSrt ML . stored

22.58C, 12 h

05r19r98, 15.98C

y78C, 48 h

12r15r98, 2.08C

y78C, 1 week

5r19r98, 15.98C

y78C, 48 h

8r05r98, 24.08C

y78C, 24 h y78C, 24 h

9r17r99, 19.08C 10r08r99, 17.58C

3.5 " 0.5 4.0 " 0.4 10.5 " 1.5 16.0 " 1.8 15.0 " 2.0 33.0 " 3.5 11.0 " 1.5 16.2 " 2.0 19.2 " 3.0 22.0 " 3.0 10.6 " 1.0 11.0 " 1.5 16.0 " 1.8

1.1 " 0.3

10r08r99, 17.58C

9.5 " 1.5 15.0 " 1.5 11.0 " 1.5 16.0 " 1.8 13.6 " 2.0 30.7 " 3.2 9.5 " 1.5 15.0 " 1.5 18.8 " 2.9 22.2 " 2.5 9.2 " 0.8 11.0 " 1.5 16.0 " 1.8

1.6 " 0.4

18C, 24 h

ML SS ML SS ML SS ML SS ML SS SS ML SS

dynamic equilibrium. Once samples are taken and enclosed in experimental tubes, this equilibrium is perturbed, and the natural difference between the microlayer and the subsurface water starts changing. Thus, our measurements of peptide hydrolysis do not reflect in-situ rates, but more the capacity of a given sample to hydrolyze peptides over a specified incubation time. The capacity for peptide hydrolysis in seawater samples was measured using fluorescent peptide substrates synthesized by condensing Lucifer Yellow Anhydride ŽLYA., a fluorescent dye, with dialanine or tetraalanine to form LYA-dialanine or LYA-tetraalanine ŽPantoja et al., 1997.. Samples were placed in polypropylene incubation bottles Žexcept on 10r29r1997 when glass was used. in the laboratory, amended with LYA-tetraalanine Žor LYA-dialanine on some occasions. to a final concentration of about 150 nM, and incubated in the dark at constant temperature Ž22.58C unless otherwise noted.. Subsamples of 0.6 ml were taken at various time intervals during the incubation. Losses of LYA-tetraalanine or LYA-dialanine and subsequent production of their fluorescent products were followed over time by high-performance liquid chromatography ŽHPLC. as described by Pantoja et al. Ž1997.. Fluorescent products were identified by comparison of retention times with synthesized standards ŽPantoja et al., 1997.. Analytical precision of the HPLC analysis was "2%. We report here means and standard

1.5 " 0.4 2.3 " 0.5 1.6 " 0.4 1.2 " 0.3 – 1.5 " 0.4

1.5 " 0.4 2.2 " 0.5 1.5 " 0.3 1.1 " 0.3 – 1.5 " 0.4

deviations of 2–4 replicate measurements of hydrolysis rate constants. Measuring peptide hydrolysis immediately was not always possible, so a reasonable sample storage method was needed. We evaluated changes in hydrolysis rates in samples stored at room temperature, at 18C, and at y78C ŽTable 1., prior to incubation. On the basis of these results Ždescribed below., water samples for hydrolysis measurements were either frozen immediately, or kept in polypropylene bottles on ice in the dark Žless than 30 min. until use. Providing a basis for quantitative comparison of hydrolysis rates is complicated when the decrease in substrate is preceded by a lag time, as we often observe for LYA peptide hydrolysis. Lag times in biological systems are common but poorly understood. Duration of a lag time can be affected by many factors, e.g., species composition, chemical characteristics of the medium, temperature and temperature changes ŽStanier et al., 1986; Vinogradov, 1977.. In this study, the biological and chemical compositions of ML and SS water differ and change seasonally. In addition, before our measurements of peptide hydrolysis, samples undergo a temperature change from in-situ to lab incubation conditions. If hydrolysis is described as a simple first-order exponential reaction as shown in curve A of Fig. 1, the hydrolysis constant, k, equals 1rt . t is defined as the time when the initial substrate concentration, C0 ,

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Table 2 Turnover times t of LYA-tetraalanine hydrolysis in microlayer ŽML. and subsurface ŽSS. water samples incubated at 22.58C, ambient bacteria concentrations ŽN., THAA concentrations and water and air temperatures for each sampling date. Samples that were frozen prior to experiments are marked with an asterisk Date

Waterrair temperature, 8C

Source

N Ž=10 5 cellsrml.

THAA ŽmM.

t, h

10r29r97

12.0r11.0

ML SS ML SS ML SS ML SS ML SS ML SS ML SS ML SS ML SS ML SS ML SS ML SS ML SS ML SS ML SS

11.5 10.3 8.9 6.8 7.5 3.8 12.5 9.6 64.4 16.8 22.3 12.6 22.5 19.1 73.6 64.5 95.6 24.3 22.9 18.5 15.0 12.3 11.9 9.7 13.1 10.6 17.4 15.2 9.5 5.5

115.0 2.4 – – – – – – 67.2 3.1 13.7 6.2 – – 19.3 2.0 7.5 1.1 4.6 1.0 12.0 0.4 5.4 0.4 5.8 0.7 29.8 0.7 30.0 0.6

13.7 31.3 13.6 30.7 14.0 27.0 12.0 16.0 12.0 17.1 11.7 13.4 9.5 15.0 8.7 11.1 18.8 22.2 11.6 18.8 15.6 24.0 11.3 17.9 11.0 31.4 12.5 24.5 15.6 26.0

12r15r97

2.0ry 2.5

2r10r98 )

4.2r4.0

3r24r98

4.5r2.0

4r8r98 )

8.2r4.2

4r21r98 )

11.5r15.0

5r19r98

15.9r23.0

7r1r98 )

20.0r18.2

8r5r98

24.0r25.2

9r18r98

21.5r14.0

10r16r98

16.3r13.0

10r30r98

13.0r7.0

11r30r98

10.7r12.0

12r7r98

12.2r18.0

2r15r99

2.3r0.0

is reduced by a factor of e Ž2.72., to Ct , where Ct s C0re. t is a mean residence time, or turnover time, and represents the time required for the initial concentration of substrate to totally disappear at a constant decay rate C0rt , the initial rate of degradation Žillustrated by curve B.. However, the commonly observed case for loss of substrate in our experiments included a lag time Žcurve C in Fig. 1.. Although LYA-tetraalanine did not decrease exponentially over the whole incubation time, hydrolysis patterns were generally similar for all samples and treatments used. So to provide a basis for comparison we chose to include the lag time with the calculation of t . Thus, we define t in this study as

the time required to reduce C0 to C0re as shown in curve C. 2.3. Ancillary time-series measurements Ambient water and air temperatures were measured at each sampling time during the field study. Bacteria were enumerated using the epifluorescence method of Hobbie et al. Ž1977. as modified by Watson et al. Ž1977.. Duplicate slides and a minimum of 100 cells per slide were counted for each sample. Counting precision was "5%. THAA concentrations were measured in both SS and ML samples at most sampling dates using vapor phase hydro-

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2.5. Controls on peptide hydrolysis

Fig. 1. ŽA. First-order rate of substrate loss, C Ž t . sC0 expŽy kt ., where C0 is the initial substrate concentration, k is the first-order rate constant, and t is elapsed incubation time. At time ta , the initial substrate concentration, C0 , would be reduced to Ct , where Ct sC0 re; ŽB. Decrease in substrate when loss is at a constant rate C0 rt ; ŽC. Exponential decrease in substrate after a lag time. In this case, t b includes the lag time.

lysis ŽTsugita et al., 1987; PICO-TAG Work Station Manual, 1987; Keil and Kirchman, 1991. and HPLC analysis as described in Mopper and Lindroth Ž1982.. Comparison of THAA between filtered samples measured either immediately after collection or after freezing for up to 3 weeks showed no significant differences. 2.4. Peptide hydrolysis in a newly-formed microlayer Subsurface water is thought to be the main source of material enriching the overlying surface microlayer ŽLiss and Duce, 1997.. Rapid film reformation results when original microlayers are removed from the sea surface, and the composition of the original and new films appears to be similar ŽWilliams et al., 1986.. To test the effect of film reformation on peptide hydrolysis, we homogenized a 50-ml SS sample collected on 12r15r1997, and allowed it to sit for 30 min in a glass beaker. During this time a new microlayer formed. The sample was then separated into two parts by withdrawing the lower half of the sample with a syringe placed at the bottom of the beaker; the upper half Žincluding the newly-formed microlayer. remained in the beaker. LYA-tetraalanine hydrolysis was then measured in each sample.

Several factors that might affect hydrolysis rate measurements were also investigated: temperature, bacteria concentration, and dissolved organic matter ŽDOM. as THAA. Since samples were incubated at 22.58C rather than at in-situ collection temperatures, ambient seawater temperatures were always lower than the incubation temperature except in August 1998. To determine how much temperature affected LYA-tetraalanine hydrolysis, samples collected on 12r15r1997 and 12r7r1999 were incubated at 28C, 108C and 22.58C and hydrolysis rate constants compared. Bacteria are believed to be the main mediators of enzymatic hydrolysis of proteinaceous matter in seawater, so the number of bacterial cells was quantified in all samples collected. We also measured changes in bacterial concentration during the incubation period on several dates Ž2r10r1998, 10r30r1998, 2r15r1999.. Extracellular Ži.e., located outside the cytoplasm. hydrolytic enzymes can remain associated with their producers so that they are part of the particulate phase, or they may be released into seawater where they can either be adsorbed onto particles or remain free in solution ŽChrost, ´ 1994.. To determine whether extracellular hydrolytic activity was in the dissolved or particulate phase, hydrolysis was measured in whole and filtered Ž0.2 mm Nuclepore. samples collected on 3r24r1998. This filtration step removes the major fraction of aquatic bacteria ŽRheinheimer, 1992.. Bacterial abundance in our filtered samples never exceeded 3% of that in unfiltered samples. Peptide hydrolysis rates may be affected by the concentration of both dissolved organic and inorganic components. DOM other than protein also contributes to bacterial growth, which in turn can affect the rate of peptide hydrolysis. Dissolved organic and inorganic compound concentrations and compositions in the microlayer and subsurface water are usually different. To investigate the effect of this dissolved material on peptide hydrolysis, we measured LYA-tetraalanine hydrolysis in SS and ML samples with the same number and type of bacteria present and at the same temperature Ž22.58C.. To equalize the number of bacteria in ML and SS water, 6-ml samples of each were collected on 3r24r1998

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and filtered through 0.2-mm sterile filters; each sample was then inoculated with 2 ml of unfiltered subsurface water. Thus, each sample retained 3r4 of its original soluble Ž- 0.2 mm. chemical components, but now had the same initial number and type of bacteria. Hydrolysis of added LYA-tetraalanine was then measured over time.

3. Results 3.1. Peptide hydrolysis patterns LYA-tetraalanine hydrolyzes over time to produce LYA-trialanine and LYA-dialanine as illustrated in microlayer and subsurface water samples from Stony Brook Harbor ŽFig. 2.. This example of hydrolysis in ML and SS samples collected 4r8r1998 is typical of all the measurements made in this study. A t-test on paired t-values collected during the entire study ŽTables 1 and 2. shows that turnover times in ML and SS waters are statistically distinct at p - 0.05. LYA-peptide hydrolysis was always faster in the sea surface microlayer ŽML. than in subsurface water ŽSS.. Although transient in both environments, LYA-trialanine concentration in the microlayer often reached higher values than in SS samples. LYA-dialanine hydrolysis measured on randomly chosen occasions, 10r29r1997 and 12r7r1998 Ždata not shown., was about 20 times slower than hydrolysis of the longer peptides, as previously observed by Pantoja et al. Ž1997.. Relative rates of hydrolysis of LYA-peptides of different chain length in both ML and SS samples were generally similar to those observed previously in bulk seawater by Pantoja and Lee Ž1999.. They occasionally observed a lag time in hydrolysis of LYA-tetra- and dipeptides in the more eutrophic waters they studied, but it was not a general characteristic. A lag time before hydrolysis began was observed for LYA-tetraalanine Ž5–12 h. and LYA-dialanine Ž150–400 h. in all our samples. The lag time varied in length and was usually shorter in ML water than in SS water, and somewhat shorter in summer than in winter. We estimated lag times for all samples collected during the time-series study from curves of the loss of added LYA peptides over time as described previously. As the sampling interval during incubations

Fig. 2. Loss of LYA-tetraalanine Ža. with time and subsequent production of LYA-trialanine Žb. and LYA-dialanine Žc. in microlayer ŽB. and subsurface Ž`. water samples collected on 4r8r1998 from Stony Brook Harbor.

was routinely 4–7 h, our lag time estimates could be in error by 2–3.5 h, or up to 40% of the estimated lag time duration. Values of t for ML and SS samples varied significantly; however, the lag time correlated Ž r 2 s 0.71. with t at the 0.95 confidence level over the entire range of values measured ŽFig. 3a.. The proportion of t made up by the lag time Žlagrt . was independent of the difference between in situ and incubation temperatures ŽFig. 3b.. We did not include data from 12r7r1998, when no lag time was observed, in calculating the correlation coefficient.

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325

temperature was unacceptable for this study. Hydrolysis Žas mean t values. in samples collected on 10r8r1999 and refrigerated for 24 h was not significantly different at the 0.05 level Ž t-test. in fresh and stored samples. Freezing and thawing also did not significantly Ž0.05 level; t-test. affect hydrolysis turnover times for samples collected at different temperatures in different seasons. Time-series samples that were frozen are indicated in Table 2. We find refrigeration and freezing acceptable for our purposes, but they should be tested for other uses. Possible artifacts that could be caused by freezing include rupturing microbial cells and releasing intracellular enzymes and THAA into the dissolved phase. During refrigeration, microbial adaptation to lower temperature could occur with consequent changes in hydrolytic processes. However, we did not observe any measurable change in t due to such processes. 3.3. Hydrolysis rate constants

Fig. 3. Relationship Ž r 2 s 0.71. between t and corresponding lag time Ža., relationship Ž r 2 s 0.0003. between lagrt and the temperature change Ž DT . from in-situ to incubation temperatures Žb.. The two points in parentheses were not considered in correlation coefficient computations.

3.2. Sample storage Refrigerating samples for up to 24 h, or freezing for up to a week were chosen as acceptable ways to preserve the natural difference between the ML and SS water. Hydrolysis in samples stored at room temperature Ž22.58C. for 12 h was about three times faster than in corresponding fresh samples collected on 5r19r1998, and the ratio of turnover times was altered ŽTable 1.. Thus, measuring hydrolysis rates in ML and SS samples that were stored at room

Values of t for LYA-hydrolysis, along with corresponding values of ambient water and air temperatures, THAA concentration, and number of bacteria for each sampling date are shown in Table 2. Mean relative deviations are as follows: "19% for t values, "5% for bacteria numbers in both ML and SS samples, and "10% and "30% for ML and SS THAA concentrations, respectively. Ambient air and water temperatures varied significantly over the year with a range of about 258. The turnover time for peptide hydrolysis varied between 9 and 31 h, and was generally shorter in samples collected in spring and summer than in fall and winter. THAA concentration was measured at 11 out of 15 sampling times. Over the year, there was no direct correlation between ambient THAA concentration and the hydrolysis rate constant Ž1rt . in either microlayer or subsurface water Ž r 2 - 0.36 for each.. However, THAA enrichment in the microlayer ŽTHAA ML r THAA SS . correlated Ž r 2 s 0.64. with the ratio of turnover times, t SS rt ML , at the 0.95 confidence level ŽFig. 4.. There was no direct correlation between the ambient bacterial enrichment of the microlayer, NML rNSS , and the ratio of turnover times, t SS rt ML , Ž r 2 s 0.20., where N is the concentration of bacteria in the samples.

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3.4. Hydrolysis in new microlayer

Fig. 4. t SS rt ML ŽRSD"38%. vs. the enrichment of THAA in the microlayer relative to subsurface water ŽRSD"43%.. The point in parenthesis was not considered in the calculation.

Turnover times presented in Table 2 were collected at a constant incubation temperature but can be extrapolated to natural conditions considering the effect of temperature on hydrolysis rate. The rate of LYA-tetraalanine hydrolysis in ML and SS samples increases with temperature. Regarding samples collected on 12r15r1997, t was 75 " 11 h in ML and 160 " 24 h in SS samples at 28C, 41 " 4 h in ML and 85 " 10 h in SS samples at 108C, and 13.6 " 2.0 h in ML and 30.7 " 4.6 h in SS samples at 22.58C. This increase in rate constants Žas 1rt . roughly corresponds to that expected from the Arrhenius relationship between chemical reaction rate and temperature. Note that the ratio t SS rt ML Ž2.1 " 0.4. did not change with temperature. The same experiment performed on samples collected 12r7r1998, when ambient parameters were considerably different from those the previous year ŽTable 2., yielded the same results Ždata not shown.. An extrapolation was made to the data collected at 22.58C assuming that 1rt doubled for every 108 rise in temperature between 28C and 22.58C. Rate constants Ž1rt . thus estimated show dramatically higher values during spring and summer ŽFig. 5a.. This figure also shows that estimated rate constants in ML and SS samples generally tracked each other over the year. However, the difference between activity in the microlayer and subsurface waters was much greater in fall and winter ŽFig. 5b.; this ratio is independent of the temperature correction.

To confirm the differences in rates between surface film and underlying water and to determine whether bulk water was the source of material enriching the microlayer and affecting the rate of peptide hydrolysis there, we measured hydrolysis in SS samples that were homogenized and then allowed to form new microlayers ŽFig. 6.. As before, LYA-tetraalanine decreased faster Žand LYA-trialanine and LYA-dialanine increased faster. in ML samples than in SS samples. Hydrolysis rates decreased in the order: original ML ) newly-formed microlayer from former SS ) original SS ) subsurface water from former SS. Thus, a newly-formed microlayer with increased enzymatic activity was established within 30 min of homogenizing SS samples, although the

Fig. 5. Ža. The annual variation in LYA-tetraalanine hydrolysis rate constant Ž1rt . in microlayer ŽB. and subsurface Ž`. water samples as corrected for temperature ŽRSD"19%.. Žb. Annual variation in the ratio t SS rt ML ŽRSD"31%..

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327

activity was not as great as in the original ML samples. 3.5. Effect of microorganisms and dissolÕed material on peptide hydrolysis Microorganisms larger than 0.2 mm are actively involved in promoting extracellular peptide hydrolysis in both ML and SS samples. Despite complete hydrolysis of LYA-tetraalanine in unfiltered samples within 1 day Žt ML s 12.0 " 1.6 h; t SS s 16.0 " 1.8 h., less than 2% of added substrate was hydrolyzed in 0.2-mm filtered samples. During incubations of ML and SS samples collected on 2r10r1998, 10r30r1998, and 2r15r1999, bacterial numbers increased somewhat with time in SS samples, but much faster Ž2–6 = . in ML water. Fig. 7 illustrates bacterial growth during the 2r15r1999 incubation. There was a f 10-h lag-time in bacterial growth in both ML and SS water samples, with the number of bacteria in ML samples increasing to almost 20 times original values before eventually reaching a plateau. To determine the effect of dissolved organic and inorganic matter on the hydrolysis rate, we filtered the ML and SS samples and then inoculated them with the same number and type of bacteria from whole SS water. The turnover

Fig. 7. Bacterial growth curve in microlayer ŽB. and subsurface Ž`. water samples during incubation at 228C with LYA-tetraalanine added to each sample to a final concentration of 142 nM. Water was sampled on 2r15r99.

time Žt . of LYA-tetraalanine in incubations containing ML samples plus the bacterial inoculate was 18 " 1.8 h. In corresponding SS samples plus bacterial inoculate, t was 24.7 " 3.0 h. For comparison, t was 12.0 " 2.3 h in unfiltered ML samples taken at the same time, and 16.0 " 3.0 h in unfiltered SS samples. Thus, although turnover times increased after filtration, the ratio t SS rt ML was about the same Žwithin experimental error. in filtered and unfiltered water. An identical experiment with samples collected in fall Ž10r30r1998. gave similar results. Bacteria always grew 3–6 times faster in ML compared to SS water in such experiments.

4. Discussion 4.1. Difference in peptide hydrolysis between ML and SS

Fig. 6. Hydrolysis of LYA-tetraalanine in original microlayer ŽB. and original subsurface Ž`. water samples collected on 3r24r98, and in subsurface water samples that were homogenized, allowed to sit and develop new microlayers, and then separated into subsamples of new subsurface water Ž,., and subsamples containing the new microlayer Žn..

We show here that differences in the biochemical environment existing between microlayer and subsurface waters can lead to differences in the rate constant but not the general pattern of hydrolysis of proteinaceous matter. We recognize that these rate factors do not represent in-situ conditions because sampling the microlayer halts the dynamic equilibrium that exists between the microlayer and subsurface waters, as mentioned earlier. However, they do indicate inherently different conditions in ML and

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SS water at the time of sampling Ži.e., capacity of the sample to hydrolyze peptides. and are thus a useful tool to investigate microlayer processes. Given that, LYA-peptide hydrolysis rate constants in the surface microlayer were consistently higher than in bulk water ŽTable 2., although the pattern of production of transient concentrations of LYA-trialanine, and then LYA-dialanine, were similar in both environments. Differences in peptide hydrolysis rates are due to differences in both concentration of substrate Žsubstrate here includes the LYA-derivatives added plus naturally occurring dissolved proteinaceous organic matter. and the numbers and types of bacteria present. Phyto-, zoo- and bacterioneuston all have different species compositions in the microlayer compared to subsurface waters ŽHardy, 1997.. Our measurements show enrichment factors for THAA in the microlayer relative to subsurface of 2–50 for Stony Brook Harbor. These enrichment factors are generally similar to measurements made by Carlucci et al. Ž1992.. Our results suggest that the influence on peptide hydrolysis of substrate concentration is more important than bacteria number or type. We measured LYA-tetraalanine hydrolysis in filter-sterilized ML and SS water samples that had identical bacterial inoculates added to each. LYA-tetraalanine hydrolysis rates were lower in filtered, inoculated samples than in whole, unfiltered samples. Since substrate concentrations were similar in each case, lower rates must have been due to fewer bacterial numbers in each case. During incubation with LYA-tetraalanine, bacteria grew faster in filtered ML samples than in filtered SS samples most likely because of the higher concentrations of THAA present there, as THAA are a major energy source for marine bacteria and serve as a good substrate for bacterial growth ŽKeil and Kirchman, 1992.. Even though hydrolysis rate constants were lower in filtered, inoculated relative to whole samples, and bacterial growth rates were greater in ML than in SS samples, the ratio of ML to SS rate constants was similar in filtered and whole samples. This suggests that differences in the amount of dissolved substrate played a more important role in establishing the difference between rates of peptide hydrolysis in ML and SS water than differences in corresponding initial microbial number or composition.

Our results on peptide hydrolysis in surface films that were newly formed from bulk water samples confirm the faster peptide hydrolysis observed in the surface microlayer, and demonstrate that material from bulk seawater can be a source of enriched material found in the microlayer. Thirty minutes after homogenization of bulk water samples in a laboratory beaker, a functional microlayer whose enrichment was sufficient to affect the peptide hydrolysis rate constant was reestablished. The higher hydrolysis rate in the subsample containing the newly-formed microlayer implies that there was substantial fractionation of the most labile substrate between the newly formed microlayer and the bulk water, since the new microlayer was a small portion of the subsample by volume. A number of factors lead to in-situ film formation ŽLiss and Duce, 1997.. High surface tension of the air–water interface energetically favors adsorption of amphiphilic organic compounds. Constituents that are scavenged from bulk water by air bubbles can accumulate at the surface. Other important factors in film formation are atmospheric deposition, wind stress, heat and mass transfer across the air–water interface, molecular diffusion and thermodiffusion, turbulent mixing of elements between microlayer and bulk water, settling of particles within the water column, solar radiation, and biochemical transformation within the layer. These processes can concentrate constituents in the microlayer at some times, and reduce the enrichment at other times. There was no wind or sunlight in our laboratory experiments, and consequently insignificant mass and energy flux from the water; deposition from the air was limited, and there was not enough time for significant biochemical change to occur. Thus, among the different mechanisms of microlayer enrichment, only the influence of surface tension seems to be significant under the conditions of our experiment. Surface tension causes stabilization of dissolved surfactants and particles on air–water interfaces Žboth new microlayer and bubble surfaces., which results in surfactant diffusion from the bulk water. Air bubbles efficiently increase the surface area, and being buoyant, actively transport the material they adsorb to the microlayer. Surface tension may thus be responsible to a large degree for the special nature of the microlayer as a habitat in the marine environment.

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4.2. Sources of enzymatic actiÕity in bulk seawater This study sheds light on two general aspects of peptide hydrolysis in bulk seawater, i.e., the source of enzymatic activity and lag times observed before LYA-tetraalanine hydrolysis commenced. Hollibaugh and Azam Ž1983. showed that most protein degrading activity in seawater is in the 0.2- to 1.0-mm size fraction, which generally includes bacterial activity. In all of our samples hydrolysis was negligible in 0.2-mm filtrates, suggesting that exoenzymes were most likely associated with their microbial producers. However, Pantoja and Lee Ž1999., studying extracellular enzymatic hydrolysis of water from Flax Pond, NY, an enclosed salt-marsh pond, demonstrated that peptidases can be found in the - 0.2-mm fraction in summer. There are several possible explanations for the differences in distribution of free and attached extracellular hydrolytic enzymes in bulk seawater between Flax Pond ŽFP. and Stony Brook Harbor ŽSBH.. Flax Pond samples were taken from a very shallow area, so sediments might serve as an immediate source of enzyme to the water column. It is also possible that bacterial grazing, which is a potential source of dissolved enzymes, was more intensive in Flax Pond water than in water of Stony Brook Harbor at the site we sampled. Nagata and Kirchman Ž1992. suggest that during grazing heterotrophic marine flagellates release their own digestive enzymes and the incompletely digested cellular components of their bacterial prey. Bacterial fragments may retain cell-associated enzymes, and these would be found in the - 0.2-mm fraction. We did not compare microbial food webs in the two areas, but bacterial numbers on 12r7r1998 were 43% higher, and temperatures almost 48 higher, in Flax Pond than in SBH, suggesting the possibility of enhanced activity in the shallower, richer waters. Lag times in peptide hydrolysis were almost always observed during this study in Stony Brook Harbor. Stanier et al. Ž1986. describe the generalized growth curve of a bacterial culture as having a lag time, an exponential growth phase, and a stationary phase, as we observed. However, since we added little new substrate to the samples, the rapid growth of bacteria might be due to the increase in temperature to laboratory values, inadvertent removal of

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predators during sampling, or prevention of diffusion of free amino acids out of the microlayer sample. In our experiments duration of the lag phase was generally proportional to t ŽFig. 3a., suggesting that either temperature, or an inherent property that depends on temperature, controls both the lag time before hydrolysis begins and the hydrolysis rate. Acclimatization time in bacterial growth depends in part on temperature of the system before transition to a new temperature ŽVinogradov, 1977.. However, although ML and SS samples in our study underwent a temperature change from various in-situ temperatures Ž2– 248C. to the incubation temperature Ž22.58C., we did not observe any correlation between temperature and lagrt , the proportion of t made up by the lag time ŽFig. 3b.. Thus, the seasonal changes we observed are unlikely to be an artifact caused by the change from in-situ to laboratory temperature. 4.3. Relation between THAA and peptide hydrolysis in the microlayer We report hydrolysis rates for our peptide analog, LYA-tetraalanine, but not for proteinaceous matter, because we know only the concentration of total hydrolyzable amino acids, and not their distribution in peptidesrproteins nor the proportion of THAA which is labile. However, since THAA concentration was always greatly enriched and turnover times always shorter in the microlayer, the peptide hydrolysis rates are almost certainly higher in the microlayer. Higher peptide hydrolysis rates, and thus rates of free amino acid supply, in the microlayer are further supported by evidence that the uptake rate of free amino acids is higher in the microlayer than subsurface waters ŽCarlucci et al., 1985, 1986.. Our preliminary measurements indicate that glutamic acid uptake rates are also higher in the microlayer of Stony Brook Harbor Žunpublished data.. There was no direct correlation observed between hydrolysis rate constant Ž1rt ., measured at a constant temperature Ž22.58C., and THAA concentration in either ML or bulk waters over the year. This would imply that bacterial growth, which influences microbial enzyme production and thus t , is not correlated with THAA concentration, as might be expected. THAA was the only fraction of the organic matter measured; as was mentioned above, THAA

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are the major energy source for aquatic bacteria. However, THAA makes up a small fraction of the dissolved organic compounds in sea water. Many other compounds could influence enzyme production. The lack of correlation between t and THAA could also be explained if not all THAA is available for use by bacteria ŽKeil and Kirchman, 1993., and if the physical form of THAA can influence its consumption rate. The ratio of peptide turnover times Žt SS rt ML . did correlate with THAA enrichment factors ŽTHAA ML rTHAA SS ., however, suggesting that each parameter is affected similarly as seasonal changes occur within the ML and SS. THAA can be present as dissolved material, colloidal aggregates or submicron particles. Elevated concentrations of dissolved organic substances in the microlayer might provoke spontaneous assembly of gel-like particles ŽChin and Verdugo, 1999., which would provide a new microenvironment for biochemical processes. Such aggregates can either be sites of intense hydrolytic enzyme activity ŽSmith et al., 1992. or havens of protection from microbial degradation ŽJohnson and Kepkay, 1992.. Adsorption of protein to particles of different sizes can have a large effect on protein degradation rates ŽNagata and Kirchman, 1997.; this could greatly affect rates in the microlayer where particles are enriched. Bubble bursting at the water surface can produce liposomes, phospholipid vesicles with microlayer water trapped inside ŽTverdislov et al., 1992.. Uptake rates of organic matter associated with such vesicles can be different from uptake rates in the surrounding microlayer. For example, Nagata and Kirchman Ž1999. showed that proteins adsorbed to the outer surface of small vesicles, and entrapped inside or integrated within the lipid bilayer, are protected from bacterial uptake. The physical form of proteinaceous compounds enriching the microlayer and their influence on enzymatic hydrolysis should be examined more closely. 4.4. Effect of season on peptide hydrolysis Two observations result from the seasonal measurements of peptide hydrolysis in ML and corresponding SS water. First, peptide hydrolysis rate constants corrected for temperature were higher in spring and summer than in fall and winter both in

ML and SS water ŽFig. 5., generally correlating with the seasonal increase in biological activity, as might be expected. The temperature correction employed here allows us to only roughly extrapolate the data collected at 22.58C to ambient conditions; it was applied to demonstrate the general seasonal trend. Second, and more surprising, the difference between microlayer and subsurface values of t Žas t ML rt SS . was greatest in fallrwinter. This difference is independent of the temperature correction we applied, and also correlates with seasonal changes in enrichment of THAA in the microlayer. There are several interrelated reasons why both the overall enrichment and composition of the surface microlayer could undergo significant seasonal changes. First, most surfactants have both hydrophilic and hydrophobic segments and can position themselves differently at surfaces, depending on temperature and other surface conditions ŽHorne, 1969.. Higher summer water temperatures can increase surfactant solubility in water, reducing enrichment factors. Second, microlayer enrichment can vary due to seasonal variations in the intensity of energy exchange between the ocean and atmosphere, which is a function of the relationship between radiative, convective and evaporative heat fluxes. For example, redistribution of inorganic ions in the microlayer correlates with intensity of evaporation from the ocean surface ŽKaravaeva et al., 1990; Kuznetsova et al., 1992.. The enrichment of organic matter in the sea surface film likely depends on thermodynamic conditions of the surface layer as well; however little is known about these effects. Third, mixing of enriched ML water with bulk seawater, which reduces the difference in constituent concentrations between ML and SS water, is more intensive in the summer. The layer of water at the air–sea interface consists of a laminar layer through which heat and mass transfer takes place by molecular diffusion, and an intermediate layer in which diffusion changes from molecular to turbulent ŽKraus and Businger, 1994.. Although the exact thickness of the layer with special chemical and biological characteristics is still a question, it is thought to be similar to or less than the thickness of the laminar layer where turbulent shear is suppressed and significant gradients are present ŽLiss and Duce, 1997;

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Zhengbin et al., 1998.. Molecular diffusion intensifies with temperature, so DOM diffuses more easily within the laminar layer in summer than in winter. It is also possible that the swimming motion of small zooneuston Žwhich are more abundant in warmer seasons. also increases mixing of the surface layer more in summer than in winter, leading to less stratification in summer. 5. Conclusions The model peptide LYA-tetraalanine was consistently hydrolysed faster in the water of the sea surface microlayer than in the underlying water. Since hydrolysis of LYA-tetraalanine is biological in origin, we conclude, as others have, that, due to accumulation of organic and inorganic matter at the boundary between ocean and atmosphere, ML water represents a more nutritious medium for microbial growth and consequently hydrolytic activity than subsurface water. We have added a new tool to quantify these differences in biological activity between the two environments. With this new tool, we have shown that the enhancement of enzymatic activity in the microlayer varied seasonally, with the relative difference between microlayer and subsurface waters being higher in fallrwinter than in springrsummer. Changes in the enhancement of hydrolytic activity followed changes in THAA enrichment of the sea surface film Žas THAA ML rTHAA SS .. Higher enrichment factors in colder seasons then in warmer seasons may be due to seasonal variations in water temperature, thermodynamic conditions of the surface microlayer, and biological activity. We did not observe any correlation between hydrolysis rates and measured THAA concentrations in either microlayer or subsurface water. The physical nature of proteinaceous compounds in the microlayer must be examined more closely to determine whether there exist protein fractions with different lability to hydrolysis. Acknowledgements We thank J. Aller for providing help in counting bacteria, S. Pantoja and D. Montluc¸on for help with probe synthesis and analysis, and M. Mulholland, A.

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Ingalls and C. Sheridan for comments on the manuscript. Support for this research was provided by the National Science Foundation Chemical Oceanography program.

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