Benthic algal nutrient bioassays traditionally have been done by measuring periphytic algal biomass that has grown on fertilized or unfertilized patches of habitat produced by nutrient-diffusing substrata (NDS). This method requires destruction of the accumulated periphyton communities and, thus, does not allow for convenient monitoring through time. Variable fluorescence methods of estimating algal biomass and photosynthetic activity have been used in aquatic environments, but generally not over different nutrient treatments and not for a substantial duration. We evaluated the use of a pulse amplitude modulated (PAM) fluorometer for measuring algal biomass and photosynthetic activity in conjunction with NDS over several weeks in a wetland under several nutrient-addition treatments. We were able to detect a significant fluorometric response as early as 1 wk into the study with addition of both N and P. Wetland periphyton was co-limited by N and P. Dark-acclimated minimal fluorescence was highly correlated with chlorophyll a in different nutrient treatments. Our results suggest that active fluorometry is a useful method for measuring periphytic responses to nutrients and for evaluating the effect of nutrient additions on overall photosynthetic efficiency.
Benthic algal responses to nutrient enrichment often have been measured by observing biomass (quantitative chlorophyll a extraction) or community composition (microscopy and morphological taxonomy) responses in nutrient-diffusing substrata (NDS) experiments (Fairchild et al. 1985, Borchardt 1996). Comparison of algal responses under ambient and nutrient-enriched conditions can be used to detect whether algal growth in an aquatic system is nutrient-limited, which nutrient is limiting algal growth, and the extent of that limitation (Francoeur et al. 1999).
In typical NDS experiments, researchers deploy growth surfaces in a water body. Some surfaces release nutrients to create local nutrient-enriched zones, whereas other surfaces do not release nutrients and serve as controls. Benthic algae are allowed to colonize and to accumulate on the surfaces and are collected later for analysis. Such experiments can be time consuming and labor intensive. The average NDS experiment requires 2 to 8 wk to complete (Biggs and Kilroy 2000), and 3 to 4 wk of exposure typically are needed to allow sufficient time for algal growth (Francoeur et al. 1999, Francoeur 2001). In addition, the biomass, community structure, or physiological assays typically applied to such experiments require destructive sampling of algal communities on NDS. This approach greatly complicates the logistics of measuring responses through time because many additional replicates are required to account for attrition caused by destructive sampling. In experiments designed for a single sampling event, destructive sampling encourages use of longer-than-necessary incubation times because investigators do not want to err by sampling too early in the colonization and growth sequence when time might not have been sufficient for nutrient-induced differences in growth rate to cause measureable differences in algal biomass. Excessive incubation times unnecessarily delay experimental completion, prevent experimenters from measuring algal responses at fine temporal scales, and may result in algal sloughing from NDS, which can mask any response to nutrient addition.
Active fluorometry can be used to measure photosynthetic performance and, under certain conditions, microalgal biomass nondestructively (Karsten et al. 1996, Serôdio et al. 1997, Honeywill et al. 2002, Consalvey et al. 2005). This method typically is used with macrophytes (Schreiber et al. 1986, Seaton and Walker 1992, Gilmore et al. 1995), but also has been used with algae (Mauzerall 1972, Samuelsson and Öquist 1977). Various portable fluorescence methods are increasingly popular alternatives for measuring algal biomass and productivity without disturbing these communities.
The 2 main methods of fluorometry currently used for in situ measurements are fast repetition rate fluorometry (FRRF) and pulse amplitude modulated (PAM) fluorometry (Kromkamp and Forster 2003). Both methods work by first measuring the quantity of photons fluoresced from Photosystem II (PSII) under ambient light conditions (the minimal fluorescence). If measurements are made on a sample held in ambient light, minimal fluorescence is denoted as F′. If measurements are made after the sample has been held for a suitable period in complete darkness (dark-acclimated), it is denoted as Fo (Table 1). In the 2nd phase of active fluorometry, periphyton are subjected to a saturating amount of photosynthetically active radiation (PAR; typically > 6000 µmol m−2 s−1; Consalvey et al. 2005) to measure the quantity of photons fluoresced when all available photocenters are saturated. This measurement is termed the maximum fluorescence and is denoted as either Fm′ or Fm, depending on whether the sample was exposed to ambient light or was dark-acclimated. PAM uses a single pulse of light typically lasting 300 to 1200 ms (Suggett et al. 2003), whereas FRRF uses a series of alternating flashes and pauses, typically 50 to 1000 ms (Kromkamp and Forster 2003) to achieve saturation. Measurements on dark-acclimated samples can be combined to calculate photosynthetic efficiency, also known as the Genty parameter, from the formula: (Fm − Fo)/Fm (Genty et al. 1989). The Genty parameter measures the maximum possible light utilization efficiency of PSII and is denoted Fv/Fm (Kromkamp and Forster 2003, Consalvey et al. 2005). Effective light utilization efficiency (measured on samples exposed to ambient light) can be calculated in a similar manner as (Fm′ − F′)/Fm′ and is denoted Fq′/Fm′ (Consalvey et al. 2005).
Table 1.
Summary of fluorescence terms. A more detailed discussion of these terms is available in the paper by Consalvey et al. (2005). PSII = photosystem II.
No general consensus exists regarding whether PAM or FRRF is superior. Few investigators have compared these 2 methods directly, and Kromkamp and Forster (2003) concluded that PAM and FRRF were correlated, but FRRF-based Fm measurements could be ∼50% higher than corresponding PAM-based Fm measurements. Suggett et al. (2003) examined phytoplankton and concluded that the 2 methods gave comparable results, but that the more-sensitive FRRF method was better suited to open-ocean studies, whereas PAM was best suited to inland freshwaters, which typically have greater algal densities.
Light exposure of algal samples has a strong effect on measured fluorescence values and their interpretation. Investigators assume that samples held in complete darkness 10 to 15 min before measurement are dark-acclimated and that all photocenters initially will be open during measurement of Fo and Fm. The measured value of Fo is determined primarily by the amount of chlorophyll a in the sample, and Fo is correlated with chlorophyll a (Kiefer et al. 1989, Kolber and Falkowski 1993, Serôdio et al. 1997, Honeywill et al. 2002, Sylvan et al. 2007). However, factors such as species composition, sample temperature, and the thickness of the periphyton layer can alter this relationship (Serôdio et al. 1997). In contrast, investigators assume that samples measured in steady-state ambient light are light-acclimated and that some photocenters will be closed during measurement of F′ and Fm′ because of photoquenching by ambient light. Therefore, the measured value of F′ is a function of the amount of chlorophyll a and the degree of photocenter closure. Thus, F′ may not be a good measure of biomass.
PSII efficiency measurements represent the capacity for efficient light use for photosynthesis as the effective efficiency under current illumination (Fq′/Fm′) or as maximum potential efficiency (Fv/Fm) (Consalvey et al. 2005). Effective PSII efficiency measurements can be used to calculate photosynthetic electron transport. Thus, efficiency measurements are correlated with algal photosynthesis as measured by 14C uptake or O2 production, although quantitative interconversion of rates is difficult (see Barranguet and Kromkamp 2000, Glud et al. 2002, Hancke at al. 2008). 14C-based measurement of photosynthetic activity under nutrient-enriched conditions has been used as a bioassay for planktonic and benthic microalgal nutrient limitation (Hameed et al. 1999, Francoeur et al. 2003). Thus, perhaps fluorescence-based measures of photosynthetic activity could be used in a similar fashion.
The nondestructive nature of active fluorometry could be a great advantage when measuring benthic algal biomass and photosynthetic responses on NDS by facilitating both measurements early in the colonization and growth process and repeated measurements through time. Our goal was to evaluate the utility of PAM fluorometry to provide rapid and repeated in situ measurements of benthic algal biomass and photosynthetic activity in nutrient-enrichment experiments by measuring algal communities incubated on varying nutrient treatments in a freshwater wetland for 3 wk.
Methods
Study site
We conducted this study at Eastern Michigan University's Loesell Wetland in Ypsilanti, Michigan, USA (lat 42.25563°N, long 83.66071°W). This 1.62-ha freshwater wetland has no surface inflow or outflow and is surrounded by residential development enclosing a region of deciduous forest that borders a wetland pool of standing water over deep, flocculent sediment. On 29 June, 6 July, 12 July, and 22 July 2007, we filtered water samples (0.7-µm glass-fiber filters GF/F) for NO3−, NH4+, and soluble reactive P (SRP) analysis. We ran analyses on a Seal AQ-2 discrete analyzer (Seal Analytical, Hampshire, UK) according to EPA-approved manufacturer's protocols for NH4+ and NO3−/NO2− (Seal Analytical 2005). We analyzed SRP with the molybdate blue method (Lind 1985). We used a YSI 63 meter (Yellow Springs Instruments, Yellow Springs, Ohio) to measure conductivity, water temperature, and pH in situ.
NDS construction and experiment
We constructed nutrient-diffusing substrata following the methods of Francoeur et al. (2013). We used a 2% agar solution with 4 different nutrient treatments: 0.5 M N as NaNO3 (N), 0.05 M P as NaHPO4 (P), both 0.5 M N and 0.05 M P (B), or a control with no additional nutrients (C). We poured agar solutions into 75-mL acid-washed polypropylene jars. After the agar had solidified, we covered it with a nitrocellulose filter (pore size = 0.8 µm) and capped the jars with lids through which a 38.1-mm-diameter hole had been bored. We replicated each nutrient treatment 5 times. We attached 1 jar of each type to a 1-m-long piece of angle-iron with ∼25 cm between jars. We suspended the NDS in the wetland pool at a depth of 10 to 15 cm below the water surface, just above the sediment, from a boardwalk running through the wetland on 29 June 2007 and left them in the wetland for 24 d. The jars were continuously submerged, except when removed for measurement. During the course of the experiment, 2 of the P replicates and 1 of the C replicates were lost.
We made measurements on 6, 12, and 22 July 2007 with a Walz Diving PAM Fluorometer (Walz, Effeltrich, Germany). We made light-acclimated measurements on cloudless days in direct, ambient sunlight during the midday hours. We made dark-acclimated measurements immediately after light-acclimated measurements by covering jars with Al cylinders for 15 min. We added a spacer to the end of the fluorescence sensor to maintain a constant distance of 1 cm between the sensor and NDS surface. We used a black plastic hood to cover each jar, the PAM fluorometer, and the investigator before making the dark-acclimated measurement. Measurements in each light treatment consisted of the minimal fluorescence (Fo or F′), the maximum fluorescence (Fm or Fm′), and the PSII efficiency (Fv/Fm or Fq′/Fm′).
Spectrophotometric chlorophyll a analysis
Immediately after the final PAM measurements on 22 July, we cut 18-mm-diameter disks from the filter of each NDS jar and froze them for later spectrophotometric chlorophyll a analysis with hot 90% ethanol extraction and acidification to correct for phaeopigments (Biggs and Kilroy 2000). At this time, the periphyton accumulated was <0.50 mm thick, and the texture of the filter NDS substrata could be seen through the covering of attached algae.
Statistical analysis
We used repeated measures and 1-way analyses of variance (ANOVAs) with subsequent post hoc Tukey's Honestly Significant Difference tests and Spearman's rank correlations in SYSTAT 11™ (Systat Software, Chicago, Illinois) to test for differences in chlorophyll a concentration and photosynthetic activity among nutrient treatments and relationships among biomass response variables.
Results
Environmental measurements
All measured environmental variables remained relatively constant during the experiment (Table 2). Water temperature and pH were maximal on 6 July. Levels of NH4+ were below the detection limits of the methods used, but levels of NO3−/NO2− decreased from 0.238 mg/L through the course of the trial. Also, levels of SRP ranged from a low of 14.96 µg/L on 29 June to a high of 59.86 µg/L on 12 July 12.
Table 2.
Environmental data from the Loesell wetland. * indicates data are from July 20. nd = not detected, SRP = soluble reactive P.
Algal biomass and photosynthetic responses to nutrients
Repeated-measures ANOVAs of F′, Fo, Fv/Fm, and Fq′/Fm′ indicated significant differences in all fluorescence variables among nutrient treatments (all p < 0.001) and over time (all p < 0.006) and significant nutrient treatment × time interactions (all p < 0.05, except for Fo, which was not affected by the interaction, p = 0.178). One-way ANOVAs considering individual variables on each sampling date revealed that PAM fluorometry could detect significant (p = 0.003) Fo responses to the B over the C treatment within 1 wk (6 July) (Fig. 1). This response also was significant on all subsequent sampling dates (12 July, p = 0.031; 22 July, p < 0.001). The nutrient-induced biomass patterns detected by Fo were confirmed by the traditional spectrophotometric biomass assay conducted on 22 July. Chlorophyll a was significantly greater on the B than the C treatment (p < 0.001; Fig. 1). By the 2nd wk, Fv/Fm and Fq′/Fm′ were significantly greater on the B than on the C treatment (Fv/Fm: 12 July, p = 0.018; 22 July, p < 0.001; Fq′/Fm′: 12 July, p = 0.011; 22 July, p < 0.001; Fig. 2A, B). Fluorescence variables never differed between N or P treatments and C treatments.
Fig.1.
Mean (+1 SE) dark-acclimated minimum fluorescence (Fo) on each date and final spectrophotometric chlorophyll a (collected 22 July) of periphyton on nutrient-diffusing substrates (C = control, P = P added, N = N added, B = both nutrients added). Numbers indicate where B treatment was significantly different from C treatment on that date. N or P treatments were never significantly different from the C treatment.
Fig.2.
Mean (+1 SE) maximum (Fv/Fm) (A) and effective (Fq′/Fm′) (B) light utilization efficiency of photosystem II (PSII) by nutrient treatment (C = control, P = P added, N = N added, B = both nutrients added). Numbers indicate where B treatment was significantly different from C treatment at that date (p < 0.05). N and P treatments were never significantly different from the C treatment. See Table 1 for explanation of fluorescence variables.
Final (July 22) Fo measurements were highly correlated (rs = 0.846, p < 0.001; Fig. 3A) but final F′ measurements were less strongly correlated (rs = 0.696, p = 0.002; Fig. 3B) with spectrophotometric measurements of chlorophyll a. Measured values of Fo, F′, and chlorophyll a were clustered by nutrient treatment, and values generally were highest on the B treatment.
Fig.3.
Scatterplots and Spearman's rank correlation of traditional spectrophotometric chlorophyll a assay results with dark-acclimated minimal fluorescence (Fo) (A) and light-acclimated minimal fluorescence (F′) (B) of periphyton on nutrient-diffusing substrates (C = control, P = P added, N = N added, B = both nutrients added) on July 22.
Discussion
Nutrient effects on algal biomass and photosynthesis
Our results suggest that traditional spectrophotometric measurement of biomass accrual and active fluorometry-based optical measurement of biomass accrual and photosynthetic activity are equivalent in their ability to detect responses to nutrient addition. Benthic algae in the Loesell wetland displayed colimitation by N and P because biomass accrual (measured by spectrophotometric chlorophyll a assay) was stimulated by simultaneous addition of N and P but not by individual N or P treatments. The colimitation observed through spectrophotometric chlorophyll a analysis was reflected in Fo and F′. These results support the theoretical and empirical arguments that Fo is an accurate indicator of algal biomass (Serôdio et al. 1997, Honeywill et al. 2002, Kromkamp and Forster 2003) and indicate that in situ optical measurements of biomass can be used in conjunction with NDS to evaluate algal responses to nutrient addition in aquatic ecosystems. If samples are measured in steady-state ambient light, some photocenters already will be closed during the measurement of F′. F′ may not be a good measure of biomass because F′ is a function of the amount of chlorophyll a and the degree of photocenter closure because of photoquenching by ambient light.
Fo was able to detect a significant biomass response to nutrient addition after only 1 wk of growth. We do not know whether spectrophotometric analysis also would have detected a significant biomass response after only 1 wk, but the nondestructive nature of the fluorometric assay allowed us to investigate the possibility of an early biomass signal without the need to deploy multiple sets of replicate substrata. Thus, use of active fluorometry could facilitate shorter NDS experimental durations and improve the temporal resolution of their conclusions.
Very high biomass could uncouple Fo and algal community biomass because fluorescence signal from deep within a periphyton mat might be quenched by overlying algae before it reaches the fluorometer (Serôdio et al. 1997, Consalvey et al. 2005). However, this limitation is not a fatal flaw because the main utility of the active fluorescence measurements in the context of an NDS experiment is to detect a biomass signal as soon as it develops, not after the community has reached maximum biomass. Investigators have attempted to quantify the relationship between the thickness of a periphyton mat and its true minimal fluorescence value when the photoquenching associated with the deeper cells has been taken into account (Forster and Kromkamp 2004, Serôdio 2004). Forster and Kromkamp (2004) found that photoquenching can occur when a microalgal cell is as shallow as 0.10 to 0.15 mm below sediment surface. The direct applicability of this estimate to our data is uncertain because our periphyton layer did not include sediments. However, the periphyton accumulated on our NDS jars was never >0.50 mm.
Effective (p < 0.001) and maximum (p < 0.001) PSII efficiencies were significantly stimulated by nutrients. These results indicate that the responses of these variables to nutrient addition can be used as a bioassay for algal nutrient limitation, as has been done with traditional 14C-based photosynthesis measurements (Hameed et al. 1999, Francoeur et al. 2003). One would expect that elevated photosynthesis would be an important mechanism driving nutrient-induced increases in microalgal biomass. Active fluorometry assays can be superior to other assay methods because their nondestructive nature allows a manageable number of experimental substrata to be measured repeatedly throughout the entire study and because they provide the opportunity to make in situ measurements of microalgal photosynthesis on fine time scales (hourly, daily; Juneau et al. 2001). This capability encourages early measurements and leads to rapid detection of microalgal biomass and photosynthetic responses to nutrient enrichment. In contrast, destructive analyses encourage use of longer incubation to ensure that responses to nutrient enrichment will have had sufficient time to occur and reach a measurable level.
Several investigators have shown that nutrient addition can rapidly (min) alter fluorescence variables (e.g., Fv/Fm, Fm) of nutrient-stressed phytoplankton, a phenomenon known as nutrient-induced fluorescence transients (Wood and Oliver 1995, Beardall et al. 2001). Our measurements were made after prolonged exposure (d to wk) of algae to nutrient additions and, thus, do not reflect these short-term transient responses. Our experiment was similar to longer-term nutrient-addition experiments in which laboratory algal cultures (Beardall et al. 2001) and natural phytoplankton communities (Sylvan et al. 2007) displayed increased Fv/Fm after relatively long (d) exposure to nutrients. Results of a single-species chemostat experiment suggested that shifts in algal Fv/Fm reliably indicated nutrient stress under unbalanced growth conditions but not under balanced growth conditions (Parkhill et al. 2001), and Kruskopf and Flynn (2006) strongly criticized the use of Fv/Fm to infer algal nutrient status. Sylvan et al. (2007) suggested that the findings of Parkhill et al. (2001) could have been caused by individualistic responses of the diatom species used in their experiment. However, balanced nutrient limitation appears to be rare in natural algal communities (see Berman-Frank and Dubinsky 1999), probably because of the multispecies nature of natural algal communities and the temporal fluctuations of environmental conditions in the field.
Applying the NDS bioassay to wetland habitats
The NDS technique was originally developed for use in lake littoral zones (Fairchild et al. 1985). Its extensive use in lakes and streams has led to synthesis of broad patterns regarding benthic algal nutrient limitation in these habitats (Borchardt 1996, Francoeur 2001). NDS have been used much less frequently in nonlittoral wetlands, preventing similar robust synthesis of wetland benthic algal nutrient limitation. We documented N and P colimitation of benthic algal biomass accrual and photosynthesis in a small Michigan wetland. This result agrees with results of previous wetland NDS experiments, which have documented nutrient limitation and co-limitation of wetland algal biomass accrual (Goldsborough et al. 2005, Scott et al. 2005). NDS results are most directly applicable to wetland benthic algal communities growing on relatively inert substrata (e.g., rocks, large woody debris, plant litter) because macrophytes and sediments can be sources of nutrients and inorganic C for epipelic algae (Vadeboncoeur and Lodge 1998, 2000, Goldsborough et al. 2005).
Factors affecting fluorometric measurements
The amount of ambient light exposure can affect measured values of light-acclimated fluorometry variables. We do not think that light-induced variation greatly affected our ability to discern nutrient-induced patterns because we made measurements only on cloudless days within a short period of time, thereby minimizing differences in ambient light exposure between replicates. In addition, light-induced variability resulting from random fluctuations in ambient light intensity during measurements should have increased stochastic variation in the measured values of light-acclimated fluorometry variables, making it less likely for us to detect nutrient-induced patterns. We are confident that any ambient light variability did not greatly affect our results because we observed clear patterns and significant differences among treatments.
The relative proportion of cyanobacteria to eukaryotic periphyton also can affect fluorometric measurements. The PAM fluorometer in our study used red (650 nm) excitation light and captured far-red (>700 nm) emission light, thereby efficiently inducing and detecting chlorophyll a fluorescence from both eukaryotic algae and cyanophytes (Walz 1998, Simis et al. 2012). Measuring cyanobacterial fluorescence with a PAM fluorometer may lead to artifactually low Fv/Fm and high minimal fluorescence readings because the ratio of cyanobacterial PSII∶PSI differs from that of eukaryotic algae and because cyanobacteria have other photosynthetic pigments that fluoresce in the far-red range (Campbell et al. 1998, Simis et al. 2012). Cursory examination of the algal communities in our study suggests that the relative abundance of chlorophytes was somewhat greater and of cyanobacteria was slightly lower in the B than in the other treatments. Thus, some of the increased Fv/Fm response in the B nutrient treatment could have been caused by a community shift away from cyanobacteria in this treatment. However, this effect probably was small in our experiment because measured Fo values increased significantly in the B treatment, a pattern opposite to that expected if among-treatment differences were caused by reduced cyanobacterial abundance in the B treatment. The close agreement of PAM and spectrophotometric biomass analyses also suggest that any potential influence of algal community composition on the PAM results was small. Nevertheless, division-level shifts in algal community composition can be induced by nutrients from NDS (Fairchild et al. 1985, Peterson and Grimm 1992), so further studies are needed to determine the degree of uncertainty caused by analysis of mixed cyanobacterial/eukaryotic algal communities.
Acknowledgements
We thank K. Judd, P. Liggit, and anonymous referees for their editorial comments. Field assistance was provided by K. Johnson, M. Schaecher, and D. Whorley. A. Johnson assisted with all laboratory analysis and technical assistance. Access was provided by Eastern Michigan University (EMU). Funding was provided by the EMU Center for Aquatic Microbial Ecology, an EMU Faculty Research Fellowship, and an EMU Biology Department Meta Hellwig Graduate Research Fellowship.
