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Microbial Ecology https://doi.org/10.1007/s00248-018-1187-z

MICROBIOLOGY OF AQUATIC SYSTEMS

Regulation of Hydrolytic Enzyme Activity in Aquatic Microbial Communities Hosted by Carnivorous Pitcher Plants Erica B. Young 1 & Jessica Sielicki 1 & Jacob J. Grothjan 1 Received: 13 December 2017 / Accepted: 10 April 2018 # Springer Science+Business Media, LLC, part of Springer Nature 2018

Abstract Carnivorous pitcher plants Sarracenia purpurea host diverse eukaryotic and bacterial communities which aid in insect prey digestion, but little is known about the functional processes mediated by the microbial communities. This study aimed to connect pitcher community diversity with functional nutrient transformation processes, identifying bacterial taxa, and measuring regulation of hydrolytic enzyme activity in response to prey and alternative nutrient sources. Genetic analysis identified diverse bacterial taxa known to produce hydrolytic enzyme activities. Chitinase, protease, and phosphatase activities were measured using fluorometric assays. Enzyme activity in field pitchers was positively correlated with bacterial abundance, and activity was suppressed by antibiotics suggesting predominantly bacterial sources of chitinase and protease activity. Fungi, algae, and rotifers observed could also contribute enzyme activity, but fresh insect prey released minimal chitinase activity. Activity of chitinase and proteases was upregulated in response to insect additions, and phosphatase activity was suppressed by phosphate additions. Particulate organic P in prey was broken down, appearing as increasing dissolved organic and inorganic P pools within 14 days. Chitinase and protease were not significantly suppressed by availability of dissolved organic substrates, though organic C and N stimulated bacterial growth, resulting in elevated enzyme activity. This comprehensive field and experimental study show that pitcher plant microbial communities dynamically regulate hydrolytic enzyme activity, to digest prey nutrients to simpler forms, mediating biogeochemical nutrient transformations and release of nutrients for microbial and host plant uptake. Keywords Extracellular enzyme . Chitinase . Protease . Phosphatase . Microbial community . Aquatic nutrient transformations . Nitrogen . Phosphorus . Sarracenia purpurea

Introduction Carnivorous pitcher plants supplement their mineral nutrition through capture of insect prey which are digested and nutrients taken up by the plant [1]. The Northern Pitcher Plant, Sarracenia purpurea subsp. purpurea L. (hereafter S. purpurea), is found in wetland habitats in North America where waterlogging slows microbial degradation of organic

Electronic supplementary material The online version of this article (https://doi.org/10.1007/s00248-018-1187-z) contains supplementary material, which is available to authorized users. * Erica B. Young [email protected] 1

Department of Biological Sciences, University of Wisconsin-Milwaukee, 3209 N Maryland Ave, Milwaukee, WI 53211, USA

matter, resulting in nutrient-depleted soils. Carnivory provides supplementary nitrogen (N), phosphorus (P) and other mineral nutrients, improving plant growth and reproductive output. Many carnivorous plants, like the Venus’ fly trap (Dionaea muscupila) and tropical Nepenthes pitcher plants, produce digestive enzymes to degrade captured insects [2, 3]. However, S. purpurea has passive traps which do not produce significant digestive enzymes [4]. The pitcher-shaped leaves collect rainwater, recruiting a community of invertebrates and microbes which mediate digestion of insect prey. The resulting aquatic detrital food web has served as a model for food web ecologists but mostly focused on the invertebrate trophic levels [5, 6]. Much less is known about the pitcher microbial communities, which are included in food web models as a Bblack box^ [7, 8]. Organic matter inputs to the pitcher plant food web are captured insects and plant detritus which falls into the pitchers. Dead insects are initially shredded by invertebrates [8], creating organic particles with larger surface area for degradation, likely involving a range of bacteria which have been

Young E. B. et al.

identified in pitchers [9, 10]. The action of microbial digestive enzymes in pitcher plants has been acknowledged [11, 12], and these enzymes likely work on detritus to transform complex organic structures into simpler molecules, providing carbon for bacterial growth, and releasing N, P, and other nutrients for plant uptake [13, 14]. But little is known about enzyme activities present, or how they are regulated in response to nutrient resources. Preliminary studies using genetic sequencing techniques indicate significant bacterial diversity in Sarracenia communities [9, 10, 15–17], and protozoans, ciliates, fungi, algae, and other protists can also inhabit pitchers [4]; these may all contribute to prey breakdown and nutrient transformations. Individual pitchers can have distinct bacterial composition [9], suggesting variable genetic potential for biochemical nutrient transformations. However, to improve food web models, this microbial diversity needs to be better connected with functional processes in the pitcher communities. In aquatic ecosystems, organic matter hydrolysis and transformations of nutrient elements between different chemical forms are mediated by microbial enzyme activities, often as extracellular or secreted enzymes [18, 19]. Several hydrolytic enzymes are particularly important in pitcher plant detrital food webs [11]. Chitinases break down insect exoskeleton chitin, a β-N-acetylglucosamine polymer, releasing monosaccharides providing organic carbon (C) and N for microbes. Chitinases are produced by aquatic bacteria [20], fungi, and arthropods [21], and plants also produce chitinases for defense [22]. Chitinases are part of the suite of digestive enzymes produced in carnivorous plants Nepenthes and Dionaea [23, 24]. Phosphatases are also ubiquitous, cleaving phosphate from organic molecules, and are examined components of organic P transformations in aquatic ecosystems [25, 26] and in carnivorous plants [27, 28]. Proteases (peptidases) in carnivorous plants hydrolyse peptide bonds of proteins in the bodies of insect prey, releasing organic C and N [29]. Proteases are also important in aquatic nutrient transformations [30]. Proteases cleave peptide bonds at specific amino acid residues, but substrate specificities can be broad and protease activity is commonly indexed as leucine aminopeptidase activity [31]. Many hydrolytic enzymes are extracellular or secreted so are readily assayed to examine microbial responses to environmental conditions [32, 33]. Extracellular enzyme activities have been quantified in bacterial communities, phytoplankton, and soils to examine microbial functions [34, 35], but such assays have not previously been applied to these novel pitcher plant communities. Pitcher plant communities are well-established models for study of food web dynamics [8], and are emerging models for study of microbial community ecology [15, 36], and so make ideal micro-ecosystems to investigate enzymatic nutrient transformations. To contribute an important functional analysis of these communities, this study presents a combination of

field sampling and manipulative experiments to examine the microbial communities, quantitatively characterize the sources of hydrolytic enzyme activities, and examine the regulation of key enzyme activities of chitinase, phosphatase, and protease, in response to nutrient and prey availability in S. purpurea pitchers. This study focusses on three aims: 1. Examine microbial communities in pitchers and relate bacterial taxa present to hydrolytic enzyme potential. We expected to identify diverse taxa, including bacteria which can produce extracellular hydrolytic enzymes. 2. Quantify hydrolytic enzyme activity and relate to pitcher prey and bacterial abundance and metabolism in pitcher communities. We hypothesized that hydrolytic enzyme activity will vary between pitchers in relation to bacterial abundance and prey detritus within pitchers, and that antibiotics which suppress bacterial metabolism will reduce enzyme activity. 3. Examine regulation of enzyme activity in response to additions of insect prey and alternative nutrient sources. We hypothesized that addition of dissolved substrates and insect prey will result in changes in enzyme activity in pitcher communities.

Methods Pitcher Plant Sampling To examine communities and their enzyme activities in natural populations in the field, S. purpurea plants were accessed in the Cedarburg Bog (43°23.2′ N, 88°0.63′ W) wetland [37], during 2008–2009 and 2014–2015. Pitcher fluid was sampled using a sterile syringe and plastic tubing inserted into the pitcher. Mature pitchers hold 20–40 ml rainwater, which was drawn up and expelled to mix contents, then samples withdrawn, stored in sterile tubes, and transported on ice. Enzyme activity was assayed within 2 h of field collection. For longer experiments, plants were labelled, single pitchers labelled with cable-ties fastened loosely around the pitcher base. In May and June 2009, 4 pitchers on each of 10 plants were sampled to compare enzyme activity variation between pitchers within plants and between plants. Twenty field pitchers were also sampled monthly June–November 2014, with water samples used for determination of cell abundance, detritus load, enzyme activities, and P concentrations. Travel to and from the field site required > 2 h, so to facilitate higher sampling frequency and to have greater control over pitcher additions for selected experiments, plants were also maintained in a greenhouse with controlled temperature, humidity, and irradiance conditions to mimic spring-fall conditions in the wetland, and to provide as cool as possible

Regulation of Hydrolytic Enzyme Activity in Aquatic Microbial Communities Hosted by Carnivorous Pitcher...

winter dormancy period. In the greenhouse, summer conditions of temperature and humidity were controlled to 15– 20 °C, 65% relative humidity, transitioning in autumn (early November–mid December) and spring (mid March–mid May) to 12–15 °C, 65% daytime humidity and 55% at night, and for winter (December–March) temperature was 2–15 °C with 55 and 65% humidity during night and day, respectively. Irradiance in the greenhouse was ambient light with mid-day shading in high summer and some supplemental light to maintain natural photoperiod in autumn and spring. Autumn, winter, and spring maximum irradiance at the plant level was 190–270 μmol photons m−2 s −1 and increased to 650– 700 μmol photons m−2 s−1 in summer. Greenhouse plants were sampled as for field populations, but samples were assayed within 30 min of collection.

Microscopy and Bacterial Diversity Bacterial abundance in pitchers was determined using water samples collected and filtered through 80-μm mesh to remove large detritus, then fixed (0.5% glutaraldehyde, PolySciences Inc., Warrington, PA) and stored at 4 °C before enumeration of cells stained with SYBRgreen (Molecular Probes, Thermo-Fisher Scientific) using epifluorescence microscopy methods of Suttle and Fuhrman [38] except using 0.2-μm black polycarbonate filters (GE Life Sciences, Pittsburgh, USA). Pitcher plant surfaces were examined using scanning electron microscopy (SEM). Pitchers cut from plants in the field were sliced into ~ 1 cm sections and fixed in the field with formalin/95% ethanol/propionic acid/water (2:10:1:7). Sections were ethanol-dehydrated, dried in a Balzers CPD020 critical point dryer, mounted to stubs, and sputter coated (20 nm gold; Structure Probe Inc., West Chester, PA, USA) and viewed at ×50–3000 magnification (Hitachi S-570 SEM). To provide an overview of the bacterial taxon composition within pitchers using genetic analysis, water samples were collected and combined from five different pitchers, prefiltered (153 μm), and cells collected on 0.22-μm polycarbonate filters, stored at − 70 °C until DNA extraction using FAST DNA spin kit for soil (MP Biomedicals). PCR amplification used 16S rRNA gene Buniversal^ 8F and 1429R primers; amplicons were purified with a QIAGEN PCR purification kit (Qiagen Inc., Valencia, CA) and cloned into pCR2.1 vector using the TOPO® TA cloning kit (Invitrogen, Carlsbad, CA). Sequencing from the 8F and 1429R primers used an ABI Big Dye Terminator Kit on an ABI Prism 3799xi Sequencer (Applied Biosystems, Foster City, CA) as described previously [39], with sequence quality control [40] and taxon identification using the Ribosomal Database Project (RDP) bacterial taxonomy classifier [41].

Enzyme Activity Assays and Localization Water collected from pitchers was pre-filtered (153 μm mesh) to remove large detritus. The proportion of enzyme activity in the dissolved fraction versus attached to particles was examined. Across samples, dissolved enzyme activity (GF/F filtrate) was a highly variable proportion of total activity, so subsequently only total activity was assayed and reported. Enzyme activities were determined using fluorometric assays adapted for black 96-well microplates [42] with the average calculated for duplicate well assays for each sample. Increases in fluorescence emission were measured at 5–10 min intervals over 60– 90 min in a microplate reader (SpectraMax Gemini XS, Molecular Devices, Sunnyvale, CA). Chitinase (N-acetyl-βD-glucosaminidase) activity was assayed using 200 μl of sample and 2.64 mM 4-methylumbelliferyl N-acetyl-β- D glucosaminide substrate in 50 mM Tris-HCl pH 8.0 (SigmaAldrich, St Louis, MO, USA) with 0.1% bovine serum albumin (BSA) (Sigma-Aldrich) [43]. Phosphatase activity was assayed in 200 μl samples with 0.72 mM 4-methylumbelliferylphosphate substrate (MUP, Sigma-Aldrich) in 50 mM TrisHCl pH 7.5 with 1% (w/v) BSA [33]. Fluorescence was read at 360 nm excitation and 420 nm emission, compared with 0– 1 μM 4-methylumbelliferone (MU, Sigma-Aldrich) standards. Protease activity was measured as leucine-aminopeptidase activity in 100 μl samples with 50 mM Tris-HCl pH 7.5 buffer and 1 mM L-leucine-7-amido-4-methylcoumarin hydrochloride substrate (L-AMC, Sigma-Aldrich). Fluorescence emission at 460 and 380 nm excitation was compared with 0–10 μM 7amino-4-methylcoumarin standards [31]. Assay control wells included buffer and substrate only, and sample and buffer only. Fluorescence emissions were converted to product concentration and plotted over time to derive a linear rate of reaction or enzyme activity. Phosphatase activity was also localized in pitcher water detritus using epifluorescence microscopy-based visualization using enzyme-linked fluorescence (ELF®-97, Invitrogen, Carlsbad CA, USA) [44]. Pitcher water samples were centrifuged (5000×g, 2–5 min) and the pellet washed with 0.2 μm filtered 100 mM phosphate-buffered saline (PBS), and resuspended in ELF reagent, incubated in the dark > 30 min, prior to a PBS rinse and visualization as previously described [33].

Sources of Enzyme Activity in Pitcher Fluid To examine the bacterial contribution to pitcher enzyme activity, responses of enzyme activity to insect prey addition were measured with and without additions of broadspectrum antibiotics, in an experiment using greenhouse plants, with five replicate pitchers for each of three treatments: (1) control with no additions, (2) addition of 20 Drosophila fruit flies (4.2–6.0 mg) to pitchers, and (3) addition of flies plus antibiotics. Flies were collected from lab

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cultures (Carolina Biological Supply) and stored frozen. Antibiotics selection was based on wetland bacterial antibiotic sensitivity [45]. The 100× stock solutions of tetracycline (2 mg/ml), erythromycin (0.25 mg/ml), both in ethanol, and ampicillin (2 mg/ml), polymyxin-B (0.25 mg/ml) in water (all Thermo-Fisher Scientific), were added to pitchers. Samples were collected at 0, 1, 6, and 13 days after addition, and assayed as above. Arthropods produce chitinase for growth regulation and molting, so the potential prey contribution to pitcher community chitinase activity was examined as release of chitinase activity from insects. In a laboratory experiment, 2, 5, or 10 Drosophila flies were added to tubes of 10 ml sterile water. Flies were added whole or briefly ground with a sterile minipestle fitted into the micro-centrifuge fly storage tube, and duplicate tubes were used for each treatment. All transfers and sampling occurred within a sterile hood. The addition of five flies represented a similar fly mass/volume ratio used for greenhouse experiments on chitinase and protease responses to substrates (below). Samples were collected at intervals over 2 weeks and assayed for chitinase.

Regulation of Phosphatase Activity and P Transformation Greenhouse-maintained pitcher plants were used to examine changes in phosphatase activity in response to phosphate additions. Pitchers all had Bnatural^ levels of prey caught over preceding months with initial pitcher water phosphate concentration of < 1.15 μM. Four replicate pitchers were used for three treatments: (1) control, no additions; (2) + 20 μM K2 HPO4; and (3) + 100 μM K2HPO4. Water was sampled at 0 and 3 h, 1, 3, and 8 days for assay of phosphatase activity. Filtered subsamples (GF/F) were used for determination of soluble molybdate-reactive P (SRP) concentrations [46]. To further explore pitcher community phosphatase activity in natural pitcher communities, a field experiment was used to test the effect of additions of more diverse prey sources and an organic P source on phosphatase activity. In July 2014, 12 replicate pitchers were labelled for each of 3 treatments: (1) control, no additions; (2) 500 μM DL-α-glycerol phosphate (Sigma-Aldrich); or (3) 14–15 mg Drosophila fruit flies plus ~ 70 mg ants (Camponotus sp.) collected from an urban garden. Pitcher samples were collected immediately after additions and after 1, 3, 7, and 14 days, and assayed for phosphatase activity. On selected days, additional samples were also collected for bacterial enumeration and for determination of P content. Dissolved phosphate (SRP) was determined as above. Total dissolved P (TDP) and total P (TP) concentrations were determined [47], and particulate P (PP) and dissolved organic P (DOP) were calculated by subtraction. After 14 days, pitcher fluid was mixed, 10 ml collected and filtered onto filters (GF/ F), and dried at 60 °C for determination of detritus mass.

Throughout the experiment, plants were covered with a 43 × 43 cm plastic canopy stapled to four wooden spikes, suspended 10–15 cm above the top of the plant, to exclude rain but maintain airflow around the plants. The opening of each pitcher was stoppered with window mesh to exclude natural prey capture.

Regulation of Chitinase and Protease Activity The short-term changes of chitinase and protease activity in response to additions of insect prey and dissolved N and C substrates were examined in greenhouse plants to allow more controlled additions and frequent sampling. The experiments were designed to test the idea that if more readily usable C or C and N sources are available for microbes, chitinase and protease enzyme synthesis may be downregulated to save energy and resources. Pitchers were disinfected and rinsed [37] and filled with the 153-μm filtered pitcher water collected 2 days previously from the field, to establish a similar initial community between pitchers. Five replicate pitchers were used for each of six treatments: (1) addition of 8–12 Drosophila flies (scaled to pitcher volume), (2) flies+ 100 μM D-glucose, (3) flies+100 μM L-glutamine, (4) flies+ 100 μM L-glutamine+100 μM D-glucose, (5) 100 μM D-glucose only, and (6) 100 μM L-glutamine only. Dissolved substrates were added to pitchers as small volumes from a 50× stock solution. Treatments were added, pitcher contents mixed, then sampled over 96 h for enzyme activity assays, and subsampled for bacterial cell enumeration.

Data Analysis Variance in field pitchers enzyme activities was analyzed between and within plants using one-way ANOVA. All analyses used Sigmaplot (v12.5, Systat Software, San Jose, CA, USA). Effects of treatments were compared using one- or two-way ANOVAwith Holm-Sidak or Tukey pairwise comparisons, with transformations applied when appropriate to avoid violating test assumptions of normal distribution and equal variance. The effect of treatments on enzyme activities in pitchers sampled over time was compared using repeated measures (RM) ANOVA. Correlations between enzyme activities were examined using Pearson product moment correlation, with Bonferroni corrections. Relationships among enzyme activity, bacterial cell abundance, and detritus were analyzed using linear regression.

Results Microbial Communities in Pitchers Insect carcasses and plant detritus accumulate at the base of pitchers, and bacteria grow on organic detritus and pitcher surfaces (Fig. 1). There was more diverse and abundant bacteria


visible with SEM on inner pitcher surfaces of mature than on young pitchers (Fig. 1e, f). Green algae (Chlorophyta) were common (Fig. 1c, d), and Haematococcus, which produces a red pigment (astaxanthin) (Fig. 1c), was dominant in some field pitchers where pitcher water was strongly red colored. Algal chlorophyll was visible as red autofluorescence, and some algal cells had been grazed by resident rotifers (Fig. 2h). A range of insect prey (ants, flies) were observed along with aquatic inhabitants Chironomidae, rotifers, algae, hyphal fungi, yeast Fig. 1 Macroscopic and microscopic views of S. purpurea pitchers. a Pitcher with collected rainwater. b Longitudinal dissection of pitcher showing accumulated insect carcasses and detritus at base of pitcher. c Green alga Haematococcus in pitcher water. d Green algae and colonial cells in matrix of hyaline filaments. e, f Scanning electron microscopy images of bacteria adhering to inner surface near the base of young pitcher (e) and mature pitcher (f); scale bars in SEM images are 10 μm

cells, bacteria, and a pine pollen grain (Fig. 2). The rotifer Habrotrocha was observed, with visible trophi (Fig. 2i). Phosphatase activity localization on pitcher detritus showed extensive labelling over prey surfaces, more intensely along exoskeleton joints and outlining fine structures which provide surfaces for bacteria to grow and accumulate (Fig. 2a– f); more degraded insects had more labelling than fresher prey (Fig. 2b, f). Algae also showed cell-associated phosphatase activity, and rotifers showed labelling within internal gut

Young E. B. et al. Fig. 2 Detritus and cells from S. purpurea pitchers with paired images of light microscopy (left) and epifluorescence visualization of phosphatase activity localization (right). a–f captured insect prey being degraded. Arrow in c indicates pine pollen grain. g–h Rotifer in pitcher water, with free algae and many algal cells inside rotifer, both showing red chlorophyll autofluorescence. i–j Two images of the rotifer Habrotrocha with visible trophi structures and phosphatase localization to internal digestive tract structures. k-l Filamentous cells, possibly fungi, with bacteria and algal cells. Scales are as shown

structures (Fig. 2g–j), possibly due to ingestion of phosphatase-expressing cells. Labelling was also along fungal filaments and on individual cells (Fig. 2k, l).

Analysis of community DNA identified 16S rRNA sequences associated with diverse bacterial taxa. Of the 176 clones identified, dominant classes were γ-proteobacteria

Regulation of Hydrolytic Enzyme Activity in Aquatic Microbial Communities Hosted by Carnivorous Pitcher... Fig. 3 a Seasonal changes in bacterial cell abundance in pitchers sampled June–November. Bars are means + standard deviation of 20 pitchers. Cell counts showed differences over time, and different letters indicate significantly different values (p < 0.001). b Relationship and linear regression between bacterial cell counts and detritus dry mass in 20 pitchers over 6 sampling events (n = 120; r2 = 0.05; p < 0.001)

(29%), β-proteobacteria (23%), Cytophagia (15%), αproteobacteria (14%), Sphingobacteria (5.7%), Flavobacteria and Clostridia (each 2.8%), and TM7, Chloroplast, Actinobacteria, Bacteroidetes also present (Table S1). Major families (> 70% of sequences) were Coxiellaceae, Comamonadaceae, Sphingomonadaceae, Rhodospirillaceae, Pseudomonadaceae, Rhizobiaceae, Aeromonadaceae, and Clostridiaceae (Fig. S1). Rickettsiella (18.75%) and Variovorax (12.6%) were the most common genera. Taxa identified represent known aquatic, sediment, soil, and wetland bacteria and those associated with algae, insects, fungi, and plants including rhizosphere inhabitants, with known chitinase, phosphatase and protease activities, cellulose degrading functions, as well as capacity for nitrate reduction, denitrification, diazotrophy, and photosynthesis (Table S1).

Quantification of Enzyme Activity—Relationships with Bacteria and Prey The dissolved hydrolytic enzyme activity (GF/F filtrate), not associated with cells or detritus in samples collected monthly from 20 pitchers over 6 months, was a variable proportion of the total activity. Dissolved chitinase activity ranged from 1 to 80% of the total (mean 22 ± 26.6%), dissolved phosphatase activity was 3.4–67% of total (29.7 ± 19.5%), and dissolved protease activity was 50–65% of total (55 ± 4.3%). Enzyme activity was variable between samples collected from different pitchers in the field population; within a single sampling event, the ratio of highest to lowest activity was > 1000 for chitinase, ~ 1000 for phosphatase, and 600 for protease. Activities measured in 4 pitchers on each of 10 plants showed no significant difference between plants (p > 0.05, one-way ANOVA on log-transformed data), indicating that pitchers could be treated as an independent, i.e., unrelated to the plant on which the pitchers were located. Across 20 field pitchers sampled monthly, there were seasonal changes in bacterial abundance with highest counts in pitchers after July (p < 0.001, one-way ANOVA on log and square-root transformed data) (Fig. 3a), while there was no

seasonal differences in detrital load (p > 0.055). However, across 120 individual pitcher samplings, bacterial cell counts were positively related to pitcher detritus (p < 0.001, linear regression on log and square-root transformed data; Fig. 3b). In the 20 pitchers sampled monthly, and in different pitchers sampled weekly over 14 days, activities of chitinase, protease, and phosphatase were significantly positively correlated with bacterial cell counts, with similar regression relationships for pitchers sampled over 6 months vs 14 days (Fig. 4). Activities of the three hydrolytic enzymes were correlated in 20 pitchers sampled over 6 months (p < 0.0001, Pearson correlation 0.391–0.444) and in 36 plants sampled three times over 2 weeks (p < 0.0001, correlation 0.551–0.638) (Fig. S2). In greenhouse pitchers with additions of flies, 1 day after additions, there was an increase in chitinase (p < 0.025, ANOVA on log-transformed data) and protease (p < 0.005, ANOVA on log-transformed data) activity (Fig. 5). In contrast, when additions of flies were accompanied by antibiotics, there was no change in chitinase or protease activity. By day 6, there was no elevated activity relative to initial, pre-addition activity. Phosphatase activity did not significantly increase following additions, though there was a decline in activity 24 h after antibiotics were added (Fig. 5). In lab experiments, chitinase activity was released from insects added into sterile water, with activity proportional to the number of flies and higher activity when ground flies were added (Fig. S3). The peak activity was around 48 h after the addition, but maximum activity with ground flies was less than 5% of maximum activity measured in pitchers with five unground fly additions within natural microbial communities.

Regulation of Enzyme Activity In a controlled greenhouse experiment, 20 and 100 μM PO43− added to pitchers was removed or transformed over the 8 days experiment, especially within the first 24 h (Fig. 6). One hundred micromolar PO43− addition resulted in significant suppression of phosphatase activity after 8 days (p < 0.001; Fig. 6). In 20 field pitchers sampled over 6 months, most of the 6–

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Fig. 4 Relationship between hydrolytic enzyme activities and bacterial cell counts in pitches sampled in the field monthly during June– November (gray symbols) and different pitchers sampled weekly for 14 days in July (open symbols). Regression lines for the two datasets are plotted (dashed for 6 month sampling, solid for 14 days sampling). All correlations were highly significant (p < 0.0001)

500 μM total P (TP) in pitchers was as particulate P (76–95%) with soluble reactive P (< 0.2–23 μM) and DOP both less than 10% of TP. In field experiments, addition of insect prey or dissolved organic P to pitchers showed increases in P pools (Fig. 7). Glycerol-P addition increased DOP and TP, though 14 days later, the DOP and PP fractions had declined dramatically. Over 14 days, the PP fraction increased in the ants+flies treatment, and DOP and SRP fractions increased. Phosphatase activity was highly variable across pitchers, and there was no significant phosphatase activity changes (two-way RM ANOVA on log and square-root transformed data).

Fig. 5 Effect of insect addition and antibiotics on hydrolytic enzyme activity in pitcher plant water. Flies and/or antibiotics added at time zero and samples collected for enzyme assays at 0, 1, and 6 days after additions. Within each treatment, significant differences in times are indicated with asterisk. Bars are means + standard deviations of five replicate pitchers

Abundance of bacterial cells did increase significantly in all three treatments over 0–7 days (p < 0.002, two-way RM ANOVA on log and square-root transformed data), but there were no clear treatment differences. The greenhouse experiment with C and N substrate additions showed more rapid increases in chitinase (within 6 h) than protease (over 24–48 h) activity but with variable enzyme activity across pitchers (Fig. 8; all differences evaluated by two-way RM ANOVA on transformed data). By 6 h, chitinase activity in pitchers with glucose+flies was higher than just glucose (p < 0.02) and glutamine+flies (p < 0.045). By 12 h,


Fig. 6 Effects of addition of 20 or 100 μM PO43− to pitcher water on phosphatase activity. Top: Drawdown of free PO43− measured in pitchers at time 0 and at days 1, 3, and 8 after phosphate additions. Bottom: Phosphatase activity measured in pitcher water sampled at time 0, and 8 days after additions. Points or bars are means ± standard deviation of four replicate pitchers. Asterisk indicates significant decline in activity over 8 days (p < 0.001)

chitinase was significantly elevated in glucose, glutamine+ flies, and glutamine+glucose+flies treatments (p < 0.005). After 24 h, glutamine only had higher chitinase activity than glucose (p < 0.01), and flies only (p < 0.05) treatments. All treatments declined by 48 h, and there were no treatment differences by 96 h. Protease activity increased most when glucose was provided and did not increase significantly when glutamine was added. Treatment differences were only observed after 48 h, when glucose+flies pitchers had significantly higher protease activity than glutamine only (p < 0.001), glutamine+flies (p < 0.003), or glucose only (p < 0.04). At 96 h, glucose+flies had higher protease activity than all other treatments (p < 0.001–p < 0.046), and the glucose and glutamine+flies showed higher activity than glutamine only (p < 0.03). The largest increases in bacterial abundance were observed with glutamine and flies; all treatments increased from 0 to 12 h (p < 0.015) but no further increases to 24 h, and no significant differences between treatments (Fig. 8).

Fig. 7 Field experiment with addition of insect prey (+Ants+Flies), DOP (glycerol-P), and control (no additions) to pitchers sampled over 14 days. Top—P fractions in pitcher water as phosphate (SRP), DOP and PP (note different P fraction scale for +Glycerol-P treatment). Error bars are the standard deviation of TP values. Middle—response of phosphatase activity. Bottom—bacterial abundance. All values are means of measurements from 12 replicate pitchers (+ standard deviation)

Discussion Pitcher Microbial Communities Sarracenia purpurea pitchers hosted diverse inquiline communities including Habrotropha rosa which is a consistent and well-studied component of Sarracenia food webs [48]. Despite reports that algae were rare in North American Sarracenia pitchers [4], small green algae were common, supporting Chlamydomonads as dominant algae in S. purpurea and the tropical pitcher plant genus Nepenthes [49, 50]. Haematococccus observed in this study has also been reported [50]. Fungi were common, as previously reported [4]. As pitchers accumulated more prey and organic detritus over

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Fig. 8 Experiments showing changes in chitinase and protease activity and bacterial abundance following the additions of flies and/or alterative microbial substrates, glutamine (glut) or glucose (gluc) or both substrates, over 96 h. Points are means of measurements in five replicate pitchers (standard deviation)

the growing season, bacterial abundance increased (Fig. 3), with a more developed bacterial flora in more mature pitchers (Fig. 1). Darlingtonia pitchers also showed increasing bacterial community diversity with time, as pitchers accumulated prey and food web inhabitants [51]. 16S rRNA gene sequencing also indicates diverse bacterial taxa in S. purpurea pitchers (Table S1). Pitcher plant micro-ecosystems are aquatic, plantassociated and are in close proximity to wetland soils—the taxa identified include representatives from these habitats. Despite the small volume of most pitchers (10–50 ml), major bacterial groups present were similar to lake ecosystems with

dominant Proteobacteria, Cytophaga, Flavobacteria, and Bacteroidetes [52]. Some similar dominant phyla (Bacteroidetes, Proteobacteria, Firmicutes) were reported in previous studies of S. purpurea [15, 36] and in Darlingtonia pitchers [51]. In Sarracenia alata, the bacterial composition was distinct from nearby soil with dominant Enterobacteriaceae [17], whereas we determined that Enterobacteriaceae was < 2% of sequences in S. purpurea (Fig. S1). Previous surveys of S. purpurea also commonly identified the enteric Serratia [9, 53], but it was not identified in this study. Common bacterial genera between S. purpurea and Nepenthes include Pedobacteria, Janthinobacterium, Pseudomonas, Micrococcus, and Fluviicola [15, 54, 55]. In addition to soil and aquatic taxa, some bacteria identified are known to be insect-associated including in gut microbiomes— Delftia, Enterobacteriaceae, and Micrococcinaceae. Deeper, mass sequencing of S. purpurea pitcher communities (Grothjan and Young, unpubl.) shows bacterial diversity consistent with this study. Different bacteria contribute different functions to pitcher communities for organic detritus breakdown, supporting the food web (Table S1). Chitinase-producing taxa include Chitinophaga, Fluviicola, Aeromonas, Pedobacter, and abundant Variovorax and Rickettsiella, which could degrade insect exoskeletons. Chitinase activity can be from algae [56] and freshwater fungi [21]. ELF could also be used to localize chitinase activity in pitcher samples [56]. Many chitinaseproducing taxa have also cellulose-degrading potential (e.g., Pedobacter); active pitchers have a waxy protective coating, but bacteria could degrade cellulose from leaf detritus caught in pitchers, and in senescing pitcher tissue. Protease activity is common in aquatic bacteria, and in dipteran larvae guts [57] which could be released into pitchers. Protease activity was identified in functional screening of S. purpurea bacterial isolates including Aeromonas [53]. Nitrogen fixation and bacterial photosynthesis potential may be important in low prey conditions. Some taxa identified can aid nutrient acquisition in the rhizosphere (e.g., Azospirillum), but similar roles within pitchers are not known. Genetic potential for enzyme production does not equate to functional enzyme activity [58]. In these pitcher plant microbial communities, assay of enzyme activity directly indicates nutrient transformation activities, including conversion of organic P to phosphate observed in field pitchers. Release of phosphate from organic compounds by phosphatase activity can be mediated by diverse community inhabitants including bacterial taxa, and eukaryotic algae [32, 33, 59] and some bacteria identified (Table S1) have been associated with P release in plant rhizospheres. However, the lack of clear antibiotic suppression of phosphatase activity suggests that phosphatase could also be from antibiotic-insensitive cells, including algae, invertebrates, and fungi which all showed phosphatase localization (Fig. 2), though antibiotic-resistant bacteria


might also be present [60]. Rotifers produce gut phosphatase [61], but chlorophyll autofluorescence suggested that ingested algal prey might also contribute phosphatase activity. Phosphatase activity localization showed accumulations along fractures and exoskeleton joints, possibly as cells concentrate in places of organic matter breakdown. Preliminary data (Menako and Young, unpubl.) showed phosphatase genes in S. purpurea communities with sequence homology to phoX in marine bacteria [62].

Quantification of Enzyme Activity—Relationships with Bacteria and Prey S. purpurea hosts one of the most diverse communities of all pitcher plant species [4], and this study demonstrates dynamic hydrolytic enzyme activity in these communities to achieve prey breakdown and nutrient transformation. Enzyme activities in S. purpurea were within similar ranges to reports for Nepenthes, and within traps of carnivorous Utricularia [3, 27]. The variable dissolved/total enzyme activity ratio was also reported in Nepenthes, with wider activity ranges in pitcher plants than in other aquatic or soil ecosystems [3], possibly due to high and variable bacterial abundance. The ranges of bacterial abundance in S. purpurea (106–107 cells/ml) and in Nepenthes (106–108 cells/ml [3]) were higher than in productive temperate freshwaters (1–8 × 106 cells/ml [63]) and aquatic sediments (4–20 × 106 cells/ml [64]). In soils, some extracellular enzyme activities can be correlated with organic matter abundance [35]. Within S. purpurea, correlations between bacterial cells, detritus, and enzyme activity indicate that enzyme activity requires cell growth and substrates. Increasing cell abundance over the growing season may relate to interaction of prey accumulation and breakdown providing substrates, increasing bacterial diversity [10, 51], and growthpromoting higher temperatures [65]. Community composition and enzyme activity may be influenced by pH [34]. Previous measurements of pH in S. purpurea pitchers showed pH over the range of 5–8 (mean 6.7, n = 55) which could influence microbial communities [37], and which is somewhat higher than pH 2–7 in Nepenthes, where the plant host can acidify the pitcher fluid [3, 4, 29, 66]. The statistical support that enzyme activity in each pitcher is independent of plant means that individual pitchers can be used for examination of hydrolytic extracellular enzyme activity regulation. Antibiotic additions showed that chitinase and protease are predominantly from bacteria, but lab experiments with fruit flies in sterile water showed that a small portion of chitinase activity in pitchers can be released from prey, during shredding of insects by invertebrates [8], but prey chitinase activity decayed within a few days of addition. Fungi can be part of pitcher plant communities (Fig. 2k) and may contribute chitinase activity [21], though our chitinase measurements in pitcher experiments with fungal inhibitors were inconclusive.

Functional laboratory surveys of bacteria isolated from S. purpurea indicated a number of chitin-degrading bacteria and fungi [53]. Pitcher plant microbial communities rapidly induced or produced de novo hydrolytic enzyme activity in response to prey additions. Chitinase activity showed the most rapid response, but still on time scales potentially related to cell growth; antibiotic suppression also suggested that activity increases require bacterial cell growth. Clear correlations among activities of chitinase, phosphatase, and protease activity measured over several months suggest that breakdown of prey requires a suite of hydrolytic enzymes functioning simultaneously. Similar correlations between hydrolytic enzymes have been reported in soils [67]. In natural conditions after pitcher opening, pitchers will capture and accumulate prey in various states of degradation, involving combinations of hydrolytic enzyme activities during breakdown. More detailed spatial and temporal analysis of enzyme activities in pitcher communities could identify any particular sequence of enzyme mediation of prey breakdown. Bacterial composition varies between pitchers [9, 15], so enzyme activity might also vary with presence of different taxa. Invertebrate communities within the pitchers may also change with time, contributing different nutrient forms [48, 68]. Other enzymes examined in carnivorous plants may be important in prey breakdown including nucleases, proteinases with other cleavage specificities, acidic phosphatases, and other aminidases, glucosidases, and cellulases [2, 3, 11, 27, 60, 69]. Many of the bacteria identified (Table S1) have cellulose degradation enzymes, which could release carbohydrate substrates from plant detritus frequently found in pitchers.

Regulation of Enzyme Activity Experiments with additions to pitchers showed regulation of enzyme activity in response to prey and nutrient sources. Phosphatase was not clearly stimulated by prey additions over 3 days as seen with chitinase and protease, but increases in phosphatase activity over a week were suggested in field plants and microbial acid phosphatase in S. purpurea pitchers has been shown to be stimulated by prey addition over 5 days [60]. Aquatic alkaline phosphatase is typically upregulated under low phosphate availability and suppressed by phosphate supply [70], while clear activity suppression was observed over 8 days with added dissolved phosphate. However, despite uptake of 100 μM phosphate, and phosphatase suppression, significant phosphatase activity was maintained, so there may be some constitutive phosphatase expression, as observed in chronically P-limited lake ecosystems [33], and in Nepenthes pitcher communities [3]. Addition of DOP and insect prey to pitcher communities had no consistent and significant effect on phosphatase activity possibly due to very high variability between field pitchers, which may relate to

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initial microbial composition, and to physiological nutrient status or community succession [51, 71]. Rapid disappearance of phosphate and DOP from pitcher water suggests community P limitation and/or rapid uptake by plant tissues, and rapid transformation of DOP into phosphate indicates phosphatase activity. Pitcher plant bacterial communities are most likely to be limited by carbon supply, but P and N may also be limiting [71], and bacteria and algae will compete with the host plant for available P [72]. In the field P addition experiment, the conversion of insect organic P into the < 153 μm particulate fraction (includes bacteria and algal cells), and also into some DOP and phosphate over 14 days, shows rapid community transformation of organic complexes in insect prey, to more bio-available P forms to support microbial metabolism, and plant P uptake. Pitcher plants take up macronutrients and many trace elements from pitchers [1, 73], but studies on nutrient forms taken up have focused mostly on N, due to availability of stable N isotopes [13, 14]. Cedarburg Bog S. purpurea nutrient stoichiometry indicated plant N limitation [37], but P limitation is possible [74]. BLuxury^ P uptake may provide P for translocation to other pitchers [75]. We hypothesized that if readily usable C and N sources were supplied to microbes, chitinase and protease enzyme synthesis would be downregulated. But when glucose and glutamine were added, the lack of clear inhibition of chitinase or protease activity suggests a more complex activity regulation, lack of regulation, or different responses across the diverse taxa present. Fungal and bacterial chitinase activity can be upregulated in response to hydrolysable substrates [76]. There may also be interactive effects of substrate availability; with additions of flies plus glucose, a readily-useable bacterial C source, cell metabolism, and use of C may have induced more N limitation, leading to higher protease activity needed to release amino acids sources from flies. The lowest protease activity, observed in the glutamine only treatment, suggests that glutamine provided both N and C sources to the microbial community. Higher protease when flies+glucose+glutamine were added may result from higher net cell metabolism stimulated by the availability of a range of substrates preferred by different taxa. Chitinase was more rapidly regulated in response to prey than protease, suggesting that some initial chitin breakdown of prey exoskeletons may be needed before prey protein is exposed and becomes available for protease action. In conclusion, this study identified eukaryotic and bacterial taxa in S. purpurea pitcher plant’s diverse communities which produce hydrolytic enzyme activities, which are dynamically regulated in response to prey additions and nutrient availability. Variability in enzyme activities may relate to community composition, but regulation of enzyme activity was consistent with other freshwater communities, with suppression and upregulation of enzyme activities, rapid transformation and

uptake of nutrient forms. These pitcher plants have served as models for food web and microbial ecology, and can also provide micro-ecosystems for examining how complex microbial communities regulate enzyme activity to mediate biogeochemical nutrient transformations [77] to support the detrital food web and host plant. Acknowledgements Research was supported by the UWM Field Station and support from the Office of Undergraduate Research to Jessica Sielicki, Addie Skillman, Jessica Mulligan, Lauren Engen, and Amy Rymaszewski. JS was also supported by UWM DIN fellowship. Terry Bott and Anne Opseth also helped with data collection. Jamie Smith and Heather Owen carried out SEM imaging. Jessica Vanderwalle helped with genetic analysis. Paul Engevold supported greenhouse experiments. We thank anonymous reviewers for their helpful suggestions.

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