Journal of Microbiological Methods 96 (2014) 62–67
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Development and validation of an open source O2-sensitive gel for physiological profiling of soil microbial communities E.S. McLamore a,⁎, J.L. Garland b, C. Mackowiak c, A. Desaunay a, N. Garland a, P. Chaturvedi a, M. Taguchi a, K. Dreaden a, John Catechis d, J.L. Ullman a a
Agricultural & Biological Engineering, University of Florida, Gainesville, FL, USA Microbiological and Chemical Exposure Assessment Research Division, U.S. Environmental Protection Agency, Cincinnati, OH, USA Department of Soil and Water Science, University of Florida, Gainesville, FL, USA d Kennedy Space Center, Space Life Sciences Laboratory, Kennedy Space Center, USA b c
a r t i c l e
i n f o
Article history: Received 1 September 2013 Received in revised form 23 October 2013 Accepted 23 October 2013 Available online 7 November 2013 Keywords: Physiological profiling Oxygen quenched dye Substrate induced respiration Assay
a b s t r a c t Community level physiological profiling is a simple, high-throughput technique for assessing microbial community physiology. Initial methods relying on redox-dye based detection of respiration were subject to strong enrichment bias, but subsequent development of a microtiter assay using an oxygen-quenched dye reduced this bias and improved the versatility of the approach. Commercial production of the oxygen microplates recently stopped, which led to the present effort to develop and validate a system using a luminophore dye (platinum tetrakis pentafluorophenyl) immobilized at the bottom of wells within a 96 well microtiter plate. The technique was used to analyze three well-characterized Florida soils: oak saw palmetto scrub, coastal mixed hardwood, and soil from an agricultural field used to grow corn silage. Substrate induced respiration was monitored by measuring respiration rates in soils under basal conditions and comparing to soils supplemented with nitrogen and various carbon sources (mannose, casein, asparagine, coumaric acid). All data was compared to a previously available commercial assay. There were no significant differences in the maximum peak intensity or the time to peak response for all soils tested (p b 0.001, α = 0.05). The experimental assay plates can be reused on soils up to four times (based on a deviation of less than 5%), where the commercial assay should not be reused. The results indicate that the new oxygen-based bioassay is a cost effective, open source tool for functional profiling of microbial communities. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Garland and Mills (Garland and Mills, 1991) developed a rapid and inexpensive screening assay to characterize microbial communities known as community level physiological profiling (CLPP). CLPP is a microwell technique that measures the aerobic respiratory demand of soil bacteria in the absence and presence of supplemental carbon and nitrogen (Brown et al., 2009; Montecchia et al., 2011; Gomez and Garland, 2012; Lehman et al., 2013). The assay allows large sets of physiological data describing microbial communities to be obtained in a relatively short period of time (usually less than 24 h). The original CLPP method (Garland and Mills, 1991) relied on a redox dye based approach for detecting aerobic respiration in microtiter plates containing relatively high concentrations of different sole carbon ⁎ Corresponding author at: 1741 Museum Rd, Rogers Hall Bldg 474, Gainesville, FL 32616, USA. Tel.: +1 352 392 1864x105. E-mail addresses: emclamor@ufl.edu (E.S. McLamore), [email protected] (J.L. Garland), echo13@ufl.edu (C. Mackowiak), aureliendesaunay@ufl.edu (A. Desaunay), ngarland929@ufl.edu (N. Garland), pchaturvedi@ufl.edu (P. Chaturvedi), [email protected]fl.edu (M. Taguchi), kmj08@ufl.edu (K. Dreaden), [email protected] (J. Catechis), jullman@ufl.edu (J.L. Ullman). 0167-7012/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.mimet.2013.10.016
sources. While useful for rapid discrimination of microbial communities, the functional relevance of the assay was limited due to the strong enrichment bias associated with the high microbial growth level required for dye reduction. Subsequent improvements (Garland et al., 2003) based on the detection of respiration using an O2 sensitive fluorophore dye reduced the problem of enrichment bias (Garland et al., 2012). Other alternative CLPP approaches have been developed based on monitoring of CO2 production in microwells (Degens et al., 2001; Campbell et al., 2003). While useful, these assessments of metabolism require nutrient supplementation at levels 10–100 times higher than that required for O2-based assays, and are not as conducive for continuous monitoring of a large number of simultaneous assays. Additionally, Folke et al (Folke et al., 2003) found that CO2 sensors are less stable than luminescent O2 sensors. The O2 microtiter assay has been used to profile soil respiratory activity in the absence and presence of supplemental carbon and nitrogen (Brown et al., 2009; Montecchia et al., 2011; Gomez and Garland, 2012; Lehman et al., 2013). Specific examples of O2-CLPP include studies of N limitation (Lehman et al., 2013), tillage and fertilization rates (Gomez and Garland, 2012), and herbicide application (Zabaloy et al., 2010, 2012). Until recently, O2-CLPP assays utilized commercial 96 well BD©
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Biosciences oxygen sensor (BDOBS). The BDOBS sensor is based on the oxygen-quenched fluorescence of a ruthenium-based dye immobilized in the bottom of a clear, 96 well polystyrene microtiter plate. The fundamental working mechanism for this sensor assay involves excitation of the dye (valence electrons are excited for a short time), and return of the valence electrons to the ground state. Upon return of valence electrons from the excited to the ground state, a photon is emitted at a wavelength higher than the excitation wavelength (i.e., lower energy state). The length of time for the valence electrons to complete this cycle is known as the fluorescence lifetime. Most fluorometers and plate readers monitor the intensity of photons emitted at the excitation wavelength (known as “intensity mode”). Many modern optical systems also directly monitor dye lifetime as a direct indicator of fluorescence/ luminescence (known as “lifetime mode”). In the last few decades, platinum porphyrin-based O2 luminescent dyes and new optical hardware have significantly improved performance of optical O2 sensors (Amao et al., 2000). These advancements have been used to develop a new generation of luminescent O2 sensors which have enhanced sensitivity, selectivity, and durability over previous technologies using rutheniumbased intensity mode sensors (Wolfbeis, 2004; McLamore et al., 2010). Recently, BD© Biosciences discontinued development of the BDOBS microtiter assay, resulting in the need for development of a new O2-based microassay for CLPP. This paper reports on a new design for O2 microtiter assays utilizing recent advancements in luminophore chemistry (platinum-based dyes) and opto-electronics (lifetime based luminescence). CLPP of several different soils using the new Pt-based microassay was compared to CLPP produced using the BDOBS system.
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Fig. 1. Conceptual schematic of the mechanism for microwell O2 technique. (A) 96 well microtiter assay. Plates are prepared and monitored using a fluorescent plate reader (with bottom laser capability). (B) Individual microtiter well depicting the soil slurry (150 μL) and the O2-sensitive film (the film thickness is ≈0.1 mm thick). The bottom of each well contains an O2-sensitive dye (platinum tetrakis pentafluorophenyl porphyrin) and polystyrene dissolved in chloroform. Excitation (X) of the dye from a laser (wavelength = 545 nm) induces fluorescence. Emission (M) is monitored at 645 nm. The presence of O2 quenches the fluorescence, and emission signal is correlated to O2 concentration based on the Stern Volmer principle.
Copper sulfate, zinc sulfate, copper chloride, manganese (II) chloride, sodium molybdate, sodium bisulfite, ammonium sulfate, and chloroform were purchased from Fisher Scientific (Waltham, PA). Platinum (II) meso-tetra pentafluorophenyl porphine (PtTFPP) was purchased from Frontier Scientific (Logan, UT). All samples were prepared with deionized water at 18.2 Ω. Polystyrene beads, L-arginine, L-asparagine, D-mannose, D-fructose, sodium acetate, propionic acid, vanillic acid and p-coumaric acid were purchased from Sigma Aldrich (St. Louis, MO).
was used immediately or stored at room temperature in a sealed glass vial with a threaded cap for up to 1 h. Solutions stored longer than ≈1 h harden and crack as the chloroform evaporates from the cocktail. BD© Falcon 96-Well cell culture plates (untreated surface) were used to prepare O2 assay plates. Aliquots (50 μL) of the dye cocktail were added to each well using a micropipette, and the plates were immediately centrifuged at 2000 rpm for 30 min to ensure homogenous distribution of the sensing membrane on the bottom of the well and to avoid cracking. Prepared assay plates were covered in foil and stored in a drawer. A Filmetrics F-20 Thin Film Analyzer was used to measure thickness of the O2 dye film. The optical properties of the sensing membrane (absorbance and lifetime) were measured using a Hitachi UV–Vis spectrometer. A FLUOstar Omega multi-mode microplate reader was used to measure dye lifetime. Excitation filters and emission filters were purchased from FLUOstar. During initial validation tests, prepared plates were inserted into the plate reader and output recorded from each well for at least five consecutive minutes. Where noted, aliquots (200 μL) of DI water or DI water + 1 mM NaHSO3 (an oxygen scavenger) were added to wells (n ≥ 3) during abiotic analysis of the prepared plates.
2.2. O2 microtiter mode of operation
2.4. Community level physiological profiling (CLPP) assays
Each fluorophore-loaded well of the BD Falcon 96-well plate received a sample of microbial communities (e.g., soil slurry), a C substrate, and amendments (N, P, or others). The plate was then placed in a temperature controlled, bottom reading, fluorescent plate reader for continuous analysis over a 24 h period. An excitation laser (X) excited the O2-sensitive luminophore in the bottom of the well, and emission (M) was measured (Fig. 1). The emission was correlated to O2 concentration in each well based on the Stern Volmer principle. Change in O2 concentration within each well was used as a direct indicator of community substrate induced respiration.
Soils from three different landscapes and soil types were collected in triplicate: oak-saw palmetto scrub (Alaquods), coastal mixed hardwoods (Hapludalfs), and an agricultural field used to grow corn silage (Quartzipsamments). Scrub soils were collected from the National Wildlife Refuge in Merritt Island, FL (Brown et al., 2009; Garland et al., 2012). Hardwood soils were collected from a protected area at the University of Florida Whitney Laboratory for Marine Biosciences (Marineland, FL). Ag soil samples were collected from UF Dairy Research Unit, Gainesville, FL. For all soil samples, ten soil cores (15 cm deep) were sampled and combined to obtain a composite sample. All samples were stored at 4 °C until analyzed. Soil samples from field sites were homogenized according to (Garland et al., 2012). Soil suspensions were prepared by mixing 10 g of homogenized soil with 25 mL of DI water in a 50 mL BD© Falcon centrifuge tube containing 5 mL sterile glass beads (2 mm diameter). Soil samples were then mixed on a vortex mixer for 1 min and aliquots of the slurry (150 μL) added to each well with a pipette. Soil assays were placed in the multi-mode plate reader and analyzed at 20 ± 1 °C. The plate reader was set up to continuously record O2 readings for 24 h (the time for one cycle of 96 measurements was 15 min). In addition to soil samples, each plate contained wells with DI water and DI water + 100 mM NaHSO3 (in triplicate) as controls.
2. Methods 2.1. Chemicals and reagents
2.3. Preparation and characterization of sensor membrane For preparing one O2-sensitive 96 well microtiter plate, 408 mg of polystyrene beads were dissolved in 5.5 g of chloroform. The solution was mixed on a vortex mixer (Vortex Genie) until the beads were completely dissolved (typically 10–15 min). Alternatively, polystyrene beads may be dissolved in chloroform at room temperature in approximately 2 h in a sealed glass vial. PtTFPP was then dissolved into the solution (concentrations from 0.1 to 0.5% w/w) in a glass vial with a threaded cap and sealed. Vials were immediately mixed for 30 s on a vortex mixer (Scientific Industries, Inc., Bohemia, NY). The dye solution
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Fig. 2. (A) Photograph of individual wells (0.4% w/w dye concentration) in a microtiter plate. Wells should be inspected prior to use by checking for visible cracks (from the bottom of the well). (B–C) Plates that were not centrifuged immediately formed relatively thick, heterogenous films that cracked. (B) photograph of a well with a few major cracks in sensing membrane (C) photograph of a well with multitude of cracks.
CLPP assays were performed using the general approach described by Zabaloy et al (Zabaloy et al., 2008) and others. Stock solutions of carbon (D-mannose, casein, L-asparagine, or p-coumaric acid at 300 mg L−1) and nitrogen (60 mg L−1 (NH4)2SO4) were prepared. The microtiter plates were pre-filled with 40 μL (per well) of one of the C solutions, N solution, or DI water. The resulting final well concentration was 50 mg C L−1 and 10 mg N L−1. 2.5. Statistical analysis Analysis of variance (ANOVA; α = 0.05) or a student's t-test were used to test for significant differences between the developed PtTFPP O2 microassay and the established commercial O2 microassay (BDOBS). Error bars in all tests represent the standard error of the arithmetic mean (n N 3). 3. Results and discussion 3.1. Film properties The average thickness of all PtTFPP films was 92 ± 4 μm. For comparison, the average thickness of the sensing membrane measured for commercial BDOBS plates was 131 ± 2 μm. If fabricated correctly, films in the bottom of the wells should be free of cracks (Fig. 2a). Plates that were not centrifuged immediately had a significantly thicker (2181 ± 94 μm) and more heterogeneous film, and cracks appeared within 24 h after the chloroform evaporated and the polystyrene hardened (Fig. 2b, c). The measured absorption bands for the PtTFPP membrane (Fig. 3) were similar to those previously reported (Chatni et al., 2009). When excited in the Soret band (395 nm) or either of the two Q bands (508 or 541 nm), the membrane displayed strong phosphorescence at 648 nm. The Soret band is within the blue region of the visible spectra,
while the Q bands are at the lower range of the green region. The highest intensity (measured as normalized relative fluorescence units, or NRFU) was 7.8 ± 0.1. This value was measured when the plate reader chamber was filled with nitrogen gas (≈ 0 kPa O2). As predicted by the Stern Volmer equation, the emission intensity was significantly lower when the chamber was filled with air (21 kPa O2) due to luminescent quenching by O2 (NRFU = 1.9 ± 0.2). This quenching effect is due to the excited triplet ground state of O2; excitation energy is released by a combination of non-radiative heat transfer, inter-particle collisions, and vibrational relaxation. Luminescent lifetime (i.e., phase shift) measured at a wavelength of 648 nm also showed O2 quenching (data not shown). For the CLPP studies below, excitation at the upper end of the Q band (≈ 545 nm) was selected based on the lower cost of filters and associated opto-electronic hardware. A number of O2 films were prepared with different concentrations of PtTFPP (while keeping polystyrene and chloroform concentrations constant). The average increase in emission intensity (NRFU) between 0 kPa and 21 kPa for all films was 70 ± 10% (Fig. 4A). Luminescent lifetime at an emission of 648 nm showed similar trends, but was more sensitive than intensity mode at low dye concentrations (Fig. 4B). The average increase in measured phase shift in 0 kPa and 21 kPa was 86 ± 7%. This significant increase in sensitivity (relative to NRFU) was due to the noise filtering capability of the phase-sensitive detection scheme in lifetime mode (Wolfbeis, 2004; McLamore et al., 2010; Chatni et al., 2009; Roche et al., 2010; Chaturvedi et al., 2013). During preliminary experiments, response time when changing from 0% to 21% O2 (or vice versa) was measured to be b60 s for all films. To maximize performance while limiting cost of the CLPP technique, 0.4% (w/w) membranes were used for all experiments to follow. For all CLPP analysis below, intensity mode was used to assess O2 dye films. As demonstrated in Fig. 4 and in the literature (Amao et al., 2000; Wolfbeis, 2004), lifetime mode analysis is more sensitive than intensity mode fluorescence. Additionally, lifetime mode analysis is
Fig. 3. A) Absorption spectra of PtTFPP dye immobilized in the bottom of a 96 well microtiter plate. B) Luminescence spectra of the sensing membrane after excitation at 545 nm. Solid line in panel B represents data collected in a nitrogen atmosphere (0 kPa O2); dashed line represents data collected in air (21 kPa O2). The emission intensity (measured in NRFU) is lower in the presence of O2 as described by the Stern Volmer principle.
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Fig. 4. A) Measured emission intensity (NRFU) at 645 nm for films exposed to 0 kPa (nitrogen gas) and 21 kPa. B) Measured phase shift (lifetime) at 0 kPa and 21 kPa O2. Lifetime mode measurements (panel B) are more sensitive than intensity mode measurements (panel A).
3.2. Comparison of PtTFPP to BDOBS plates In a preliminary study, CLPP using PtTFPP O2 plates prepared with a dye concentration of 0.3% (w/w) showed markedly lower levels of peak response compared to the commercial BDOBS O2 assay (see supplemental data, Fig. S1). Overall, peak response appeared to be limited to ~2 NRFU in the PfTFPP plates. To resolve this problem, PtTFPP plates were prepared with a dye concentration of 0.4% (w/w) to increase the maximum emission signal and the assays were repeated. When using PtTFPP plates with a dye concentration of 0.4%, there was no statistical difference in the maximum peak intensity (NRFU) when compared to BDOBS plates for the same soil slurries (p b 0.001; α = 0.05). For agricultural soils, supplementation of nitrogen or carbon increased average maximum intensity in all soils tested. Fig. 5 shows a representative transient plot showing peak NRFU response. Addition of supplemental nitrogen (10 mg L-1 (NH4)2SO4) caused a significant increase in peak aerobic respiration (23.2 ≈ 3.4%), but did not significantly change the time to peak (p b 0.001, α = 0.05). For all soils tested, addition of nitrogen alone caused an increase in peak NRFU of approximately 50%, while addition of casein, asparagine, and coumaric acid with nitrogen caused an increase in peak NRFU of 25–30% (Fig. 6). The largest increase was after the addition of nitrogen and mannose (72%). These trends are similar to those reported by Garland et al for similar soils using BDOBS plates (Garland et al., 2012), although the peak NRFU values in these samples are higher than those previously reported. For the Scrub and Hardwood soils, supplemental carbon addition did not always result in a significant increase in peak respiration. Mannose, coumaric acid and acetate significantly increased peak NRFU response, but casein and asparagine supplementation did not significantly increase peak NRFU. Regardless of N amendment, mannose, coumaric acid and acetate had the largest impact on SIR in Hardwood and Scrub soils. In Scrub soils, the major difference was a noted increase in asparagine metabolism when nitrogen was supplemented. While these minor differences in soil physiology are still being investigated, the important finding here is that no significant
differences were measured between the experimental PtTFPP and conventional BDOBS assays. The measured time associated with peak fluorescence (i.e., lag time) from samples treated with the PtTFPP dye method was not significantly different than with the BDOBS method (p b 0.0001, α = 0.05). Together with the peak response data, the new method appears comparable to the BDOBS method for CLPP (see supplemental data, Fig. S2 for time to peak data). The PtTFPP and commercial BDOBS plates were re-used seven times to estimate the shelf life and/or any sensor hysteresis. The average drift (i.e., percent change in output) was determined by calculating the mean percent change in peak response for each well in the assay, and then calculating the arithmetic mean of all peak NRFU responses (representing the net drift in sensor output). After each experiment, plates were rinsed, and soaked in DI water at 30 °C for 6 h, and air dried in the dark for 6 h. The threshold for performance was a 5% loss in performance (noted by a horizontal dashed line in Fig. 7). Based on this threshold, the PtTFPP plates may be re-used up to four times, while the BDOBs plates should not be used more than once. The reusability of BDOBS plates reported here was similar to results reported by Birmele et al (Birmele et al., 2006) characterizing planktonic Bacillus subtilis 1A2 and Pseudomonas aeruginosa PA01 respiration. This difference in performance between the two dye films is likely due to reduced photobleaching of platinum luminophores relative to
2.4 Scrub: Acetate (no N) Scrub: Acetate + N)
2.2 2.0
Emission [NRFU]
known to cause less photobleaching than intensity mode measurements. Photobleaching is a common problem with fluorescent dyes and causes signal drift. Although lifetime mode measurements are preferred if possible, intensity mode measurements were chosen for the following reasons: 1) a wealth of data exists in the literature for intensity mode measurements with BDOBS plates, 2) plate readers with lifetime mode capability are relatively new and therefore may limit the broad applicability of the technique, and 3) the lifetime of PtTFPP is different than the that of the ruthenium dye within the BDOBS plate, potentially imparting bias on the comparative study.
1.8 1.6 1.4 1.2 1.0 0.8 0
500
1000
1500
2000
2500
3000
Time [min] Fig. 5. Representative plot showing transient response of PtTFPP assay for an oak saw palmetto scrub soil sample with supplemented acetate. The closed circles show the transient response in the absence of supplemental nitrogen (10 mg L−1 (NH4)2SO4); the open triangles show the respiratory response with supplemental acetate and nitrogen.
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Percent change in peak response [%]
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PtTFPP plates BDOBS plates
0
-5
-10
-15
-20
-25 0
1
2
3
4
5
6
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8
Number of repeated uses [#] Fig. 7. Comparison of change in PtTFPP (0.4% w/w dye concentration) and BDOBS peak response during repeated use. The horizontal dotted line represents a 5% reduction in measured peak O2 response for all wells in the assay.
ruthenium-based dyes (Wolfbeis, 2004). In addition, the PtTFPP is nonionic and hydrophobic, reducing the leaching rate from the sensing membrane. Since output from all wells (both abiotic and biotic) were grouped together before calculating the percent change in output, the data represent the combined effects of photobleaching during measurement as well as hysteresis due to exposure of the dye to organic acids commonly found in soils (e.g., malate, citrate). If available, lifetime mode is recommended for all CLPP using the PtTFPP assay due to the enhance performance and increased longevity of the sensor. Available commercial systems such as the sensor dish reader (SDR) by Presens and MitoExpress© by LuxCel Biosciences. The SDR performs a similar oxygen analysis but is only available in 6 well or 24 well format. The SDR has the advantage of simultaneous analysis of pH and O2 within each, but has some disadvantages for soil analyses. In addition to prohibitive costs for soil analyses of large fields (where thousands of samples may be required), multi-parameter systems such as the SDR require complex three dimensional calibration curves and thus technician training. The MitoExpress© instrument also measures pH and O2, but is primarily intended for analysis of suspended cells and is not reliable for rapid multiplexing with natural soils. These challenges and others are discussed in detail in the review by Chaturvedi et al (Chaturvedi et al., 2013). Table 1 provides a basic cost summary for potential users of the technique. The total cost for preparing one 96 well plate for CLPP analysis in 2013, is approximately $15 USD (not accounting for potential reuse of the plate). Ordering supplies in bulk can reduce this cost by up to 8%. Use of platinum porphyrin dyes in opto-electronics is expected to increase in the coming decade, which will likely reduce the cost of assay preparation. The open source recipe described here provides soil scientists with a reliable recipe for preparing O2-sensitive gels used for physiological profiling of soil microbial communities.
Table 1 Cost summary in 2013 (in USD) for PtTFPP plates used for CLPP analysis. The total cost per plate is approximately $15 for the recipe with 0.4% dye.
Fig. 6. Comparison of PtTFPP (black bars) and BDOBS (gray bars) peak NRFU for samples in the presence and absence of various carbon/nitrogen substrates. O2-CLPP assays for (A) Agricultural soil, (B) Scrub soil, and (C) Hardwood soil. Carbon supplement contained 300 mg L−1 of indicated compound; (+N) indicates addition of 60 mg L1 (NH4)2SO4.
Item
Company
Pt(II) meso-tetra Pentafluorophenyl porphine BD Falcon 96 well plates Chloroform Polystyrene beads
Frontier Scientific PtT975 BD Biosciences Sigma Aldrich Sigma Aldrich
Product no.
353916 472476-1 L 39411-50G-F
Cost per plate [USD] $11.24 $3.02 $0.59 $0.12
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Validation of the new recipe presented here allows for continued development of the O2-based CLPP approach. As discussed in the introduction, this technique provides a less biased method for physiological profiling compared to previous formats (i.e., Biolog plates) due to lower substrate concentrations and elimination of redox dyes and proprietary growth supporting nutrients. However, the purpose of the CLPP approach, even with the O2-based platform, is rapid screening of the physiological capacity of microbial communities and not direct assessment of in situ physiological activity. Recent work with the BDoxy plates showed that the O2 peak responses between 6–24 h correspond to less than 1 to ~4 doublings (Garland et al., 2012). Therefore, the assay is useful for determining how communities are poised to utilize different types of substrate and what types of factors (e.g., N) may be limiting activity. This is particularly important when intensive spatiotemporal sampling and/or multiple experiment spatial treatments are of interest. When estimating specific rates of activity are of concern, alternative methods should be employed.
4. Conclusion Previous researchers have verified the utility of the O2-CLPP to assess the physiological status of microbial communities, particularly as applied to soil. To meet this need, a simple recipe was developed for fabrication of an open source O2-sensitive 96 well assay. The assay screens soil community physiology within a short period (≈24 h) by monitoring aerobic respiratory activity in microtiter wells using an oxygenquenched luminescent dye. Following the CLPP approach developed with commercial assays, substrate induced respiration and endogenous respiration were monitored for until a peak respiratory response is reached. The performance of the sensor technology is equivalent to a previously available commercial assay, but the sensor developed herein proved to be more robust and may be amenable to reuse.
Acknowledgments The authors would like to thank the Florida Agriculture Experimental Station (M.D. Dukes), the UF Excellence Award, and the IFAS Early Career Award (CRIS No. 005062) for funding. The authors also thank David Julian for the assistance with plate reader analysis.
Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.mimet.2013.10.016.
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Contents lists available at ScienceDirect
Journal of Microbiological Methods journal homepage: www.elsevier.com/locate/jmicmeth
Development and validation of an open source O2-sensitive gel for physiological profiling of soil microbial communities E.S. McLamore a,⁎, J.L. Garland b, C. Mackowiak c, A. Desaunay a, N. Garland a, P. Chaturvedi a, M. Taguchi a, K. Dreaden a, John Catechis d, J.L. Ullman a a
Agricultural & Biological Engineering, University of Florida, Gainesville, FL, USA Microbiological and Chemical Exposure Assessment Research Division, U.S. Environmental Protection Agency, Cincinnati, OH, USA Department of Soil and Water Science, University of Florida, Gainesville, FL, USA d Kennedy Space Center, Space Life Sciences Laboratory, Kennedy Space Center, USA b c
a r t i c l e
i n f o
Article history: Received 1 September 2013 Received in revised form 23 October 2013 Accepted 23 October 2013 Available online 7 November 2013 Keywords: Physiological profiling Oxygen quenched dye Substrate induced respiration Assay
a b s t r a c t Community level physiological profiling is a simple, high-throughput technique for assessing microbial community physiology. Initial methods relying on redox-dye based detection of respiration were subject to strong enrichment bias, but subsequent development of a microtiter assay using an oxygen-quenched dye reduced this bias and improved the versatility of the approach. Commercial production of the oxygen microplates recently stopped, which led to the present effort to develop and validate a system using a luminophore dye (platinum tetrakis pentafluorophenyl) immobilized at the bottom of wells within a 96 well microtiter plate. The technique was used to analyze three well-characterized Florida soils: oak saw palmetto scrub, coastal mixed hardwood, and soil from an agricultural field used to grow corn silage. Substrate induced respiration was monitored by measuring respiration rates in soils under basal conditions and comparing to soils supplemented with nitrogen and various carbon sources (mannose, casein, asparagine, coumaric acid). All data was compared to a previously available commercial assay. There were no significant differences in the maximum peak intensity or the time to peak response for all soils tested (p b 0.001, α = 0.05). The experimental assay plates can be reused on soils up to four times (based on a deviation of less than 5%), where the commercial assay should not be reused. The results indicate that the new oxygen-based bioassay is a cost effective, open source tool for functional profiling of microbial communities. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Garland and Mills (Garland and Mills, 1991) developed a rapid and inexpensive screening assay to characterize microbial communities known as community level physiological profiling (CLPP). CLPP is a microwell technique that measures the aerobic respiratory demand of soil bacteria in the absence and presence of supplemental carbon and nitrogen (Brown et al., 2009; Montecchia et al., 2011; Gomez and Garland, 2012; Lehman et al., 2013). The assay allows large sets of physiological data describing microbial communities to be obtained in a relatively short period of time (usually less than 24 h). The original CLPP method (Garland and Mills, 1991) relied on a redox dye based approach for detecting aerobic respiration in microtiter plates containing relatively high concentrations of different sole carbon ⁎ Corresponding author at: 1741 Museum Rd, Rogers Hall Bldg 474, Gainesville, FL 32616, USA. Tel.: +1 352 392 1864x105. E-mail addresses: emclamor@ufl.edu (E.S. McLamore), [email protected] (J.L. Garland), echo13@ufl.edu (C. Mackowiak), aureliendesaunay@ufl.edu (A. Desaunay), ngarland929@ufl.edu (N. Garland), pchaturvedi@ufl.edu (P. Chaturvedi), [email protected]fl.edu (M. Taguchi), kmj08@ufl.edu (K. Dreaden), [email protected] (J. Catechis), jullman@ufl.edu (J.L. Ullman). 0167-7012/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.mimet.2013.10.016
sources. While useful for rapid discrimination of microbial communities, the functional relevance of the assay was limited due to the strong enrichment bias associated with the high microbial growth level required for dye reduction. Subsequent improvements (Garland et al., 2003) based on the detection of respiration using an O2 sensitive fluorophore dye reduced the problem of enrichment bias (Garland et al., 2012). Other alternative CLPP approaches have been developed based on monitoring of CO2 production in microwells (Degens et al., 2001; Campbell et al., 2003). While useful, these assessments of metabolism require nutrient supplementation at levels 10–100 times higher than that required for O2-based assays, and are not as conducive for continuous monitoring of a large number of simultaneous assays. Additionally, Folke et al (Folke et al., 2003) found that CO2 sensors are less stable than luminescent O2 sensors. The O2 microtiter assay has been used to profile soil respiratory activity in the absence and presence of supplemental carbon and nitrogen (Brown et al., 2009; Montecchia et al., 2011; Gomez and Garland, 2012; Lehman et al., 2013). Specific examples of O2-CLPP include studies of N limitation (Lehman et al., 2013), tillage and fertilization rates (Gomez and Garland, 2012), and herbicide application (Zabaloy et al., 2010, 2012). Until recently, O2-CLPP assays utilized commercial 96 well BD©
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Biosciences oxygen sensor (BDOBS). The BDOBS sensor is based on the oxygen-quenched fluorescence of a ruthenium-based dye immobilized in the bottom of a clear, 96 well polystyrene microtiter plate. The fundamental working mechanism for this sensor assay involves excitation of the dye (valence electrons are excited for a short time), and return of the valence electrons to the ground state. Upon return of valence electrons from the excited to the ground state, a photon is emitted at a wavelength higher than the excitation wavelength (i.e., lower energy state). The length of time for the valence electrons to complete this cycle is known as the fluorescence lifetime. Most fluorometers and plate readers monitor the intensity of photons emitted at the excitation wavelength (known as “intensity mode”). Many modern optical systems also directly monitor dye lifetime as a direct indicator of fluorescence/ luminescence (known as “lifetime mode”). In the last few decades, platinum porphyrin-based O2 luminescent dyes and new optical hardware have significantly improved performance of optical O2 sensors (Amao et al., 2000). These advancements have been used to develop a new generation of luminescent O2 sensors which have enhanced sensitivity, selectivity, and durability over previous technologies using rutheniumbased intensity mode sensors (Wolfbeis, 2004; McLamore et al., 2010). Recently, BD© Biosciences discontinued development of the BDOBS microtiter assay, resulting in the need for development of a new O2-based microassay for CLPP. This paper reports on a new design for O2 microtiter assays utilizing recent advancements in luminophore chemistry (platinum-based dyes) and opto-electronics (lifetime based luminescence). CLPP of several different soils using the new Pt-based microassay was compared to CLPP produced using the BDOBS system.
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Fig. 1. Conceptual schematic of the mechanism for microwell O2 technique. (A) 96 well microtiter assay. Plates are prepared and monitored using a fluorescent plate reader (with bottom laser capability). (B) Individual microtiter well depicting the soil slurry (150 μL) and the O2-sensitive film (the film thickness is ≈0.1 mm thick). The bottom of each well contains an O2-sensitive dye (platinum tetrakis pentafluorophenyl porphyrin) and polystyrene dissolved in chloroform. Excitation (X) of the dye from a laser (wavelength = 545 nm) induces fluorescence. Emission (M) is monitored at 645 nm. The presence of O2 quenches the fluorescence, and emission signal is correlated to O2 concentration based on the Stern Volmer principle.
Copper sulfate, zinc sulfate, copper chloride, manganese (II) chloride, sodium molybdate, sodium bisulfite, ammonium sulfate, and chloroform were purchased from Fisher Scientific (Waltham, PA). Platinum (II) meso-tetra pentafluorophenyl porphine (PtTFPP) was purchased from Frontier Scientific (Logan, UT). All samples were prepared with deionized water at 18.2 Ω. Polystyrene beads, L-arginine, L-asparagine, D-mannose, D-fructose, sodium acetate, propionic acid, vanillic acid and p-coumaric acid were purchased from Sigma Aldrich (St. Louis, MO).
was used immediately or stored at room temperature in a sealed glass vial with a threaded cap for up to 1 h. Solutions stored longer than ≈1 h harden and crack as the chloroform evaporates from the cocktail. BD© Falcon 96-Well cell culture plates (untreated surface) were used to prepare O2 assay plates. Aliquots (50 μL) of the dye cocktail were added to each well using a micropipette, and the plates were immediately centrifuged at 2000 rpm for 30 min to ensure homogenous distribution of the sensing membrane on the bottom of the well and to avoid cracking. Prepared assay plates were covered in foil and stored in a drawer. A Filmetrics F-20 Thin Film Analyzer was used to measure thickness of the O2 dye film. The optical properties of the sensing membrane (absorbance and lifetime) were measured using a Hitachi UV–Vis spectrometer. A FLUOstar Omega multi-mode microplate reader was used to measure dye lifetime. Excitation filters and emission filters were purchased from FLUOstar. During initial validation tests, prepared plates were inserted into the plate reader and output recorded from each well for at least five consecutive minutes. Where noted, aliquots (200 μL) of DI water or DI water + 1 mM NaHSO3 (an oxygen scavenger) were added to wells (n ≥ 3) during abiotic analysis of the prepared plates.
2.2. O2 microtiter mode of operation
2.4. Community level physiological profiling (CLPP) assays
Each fluorophore-loaded well of the BD Falcon 96-well plate received a sample of microbial communities (e.g., soil slurry), a C substrate, and amendments (N, P, or others). The plate was then placed in a temperature controlled, bottom reading, fluorescent plate reader for continuous analysis over a 24 h period. An excitation laser (X) excited the O2-sensitive luminophore in the bottom of the well, and emission (M) was measured (Fig. 1). The emission was correlated to O2 concentration in each well based on the Stern Volmer principle. Change in O2 concentration within each well was used as a direct indicator of community substrate induced respiration.
Soils from three different landscapes and soil types were collected in triplicate: oak-saw palmetto scrub (Alaquods), coastal mixed hardwoods (Hapludalfs), and an agricultural field used to grow corn silage (Quartzipsamments). Scrub soils were collected from the National Wildlife Refuge in Merritt Island, FL (Brown et al., 2009; Garland et al., 2012). Hardwood soils were collected from a protected area at the University of Florida Whitney Laboratory for Marine Biosciences (Marineland, FL). Ag soil samples were collected from UF Dairy Research Unit, Gainesville, FL. For all soil samples, ten soil cores (15 cm deep) were sampled and combined to obtain a composite sample. All samples were stored at 4 °C until analyzed. Soil samples from field sites were homogenized according to (Garland et al., 2012). Soil suspensions were prepared by mixing 10 g of homogenized soil with 25 mL of DI water in a 50 mL BD© Falcon centrifuge tube containing 5 mL sterile glass beads (2 mm diameter). Soil samples were then mixed on a vortex mixer for 1 min and aliquots of the slurry (150 μL) added to each well with a pipette. Soil assays were placed in the multi-mode plate reader and analyzed at 20 ± 1 °C. The plate reader was set up to continuously record O2 readings for 24 h (the time for one cycle of 96 measurements was 15 min). In addition to soil samples, each plate contained wells with DI water and DI water + 100 mM NaHSO3 (in triplicate) as controls.
2. Methods 2.1. Chemicals and reagents
2.3. Preparation and characterization of sensor membrane For preparing one O2-sensitive 96 well microtiter plate, 408 mg of polystyrene beads were dissolved in 5.5 g of chloroform. The solution was mixed on a vortex mixer (Vortex Genie) until the beads were completely dissolved (typically 10–15 min). Alternatively, polystyrene beads may be dissolved in chloroform at room temperature in approximately 2 h in a sealed glass vial. PtTFPP was then dissolved into the solution (concentrations from 0.1 to 0.5% w/w) in a glass vial with a threaded cap and sealed. Vials were immediately mixed for 30 s on a vortex mixer (Scientific Industries, Inc., Bohemia, NY). The dye solution
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Fig. 2. (A) Photograph of individual wells (0.4% w/w dye concentration) in a microtiter plate. Wells should be inspected prior to use by checking for visible cracks (from the bottom of the well). (B–C) Plates that were not centrifuged immediately formed relatively thick, heterogenous films that cracked. (B) photograph of a well with a few major cracks in sensing membrane (C) photograph of a well with multitude of cracks.
CLPP assays were performed using the general approach described by Zabaloy et al (Zabaloy et al., 2008) and others. Stock solutions of carbon (D-mannose, casein, L-asparagine, or p-coumaric acid at 300 mg L−1) and nitrogen (60 mg L−1 (NH4)2SO4) were prepared. The microtiter plates were pre-filled with 40 μL (per well) of one of the C solutions, N solution, or DI water. The resulting final well concentration was 50 mg C L−1 and 10 mg N L−1. 2.5. Statistical analysis Analysis of variance (ANOVA; α = 0.05) or a student's t-test were used to test for significant differences between the developed PtTFPP O2 microassay and the established commercial O2 microassay (BDOBS). Error bars in all tests represent the standard error of the arithmetic mean (n N 3). 3. Results and discussion 3.1. Film properties The average thickness of all PtTFPP films was 92 ± 4 μm. For comparison, the average thickness of the sensing membrane measured for commercial BDOBS plates was 131 ± 2 μm. If fabricated correctly, films in the bottom of the wells should be free of cracks (Fig. 2a). Plates that were not centrifuged immediately had a significantly thicker (2181 ± 94 μm) and more heterogeneous film, and cracks appeared within 24 h after the chloroform evaporated and the polystyrene hardened (Fig. 2b, c). The measured absorption bands for the PtTFPP membrane (Fig. 3) were similar to those previously reported (Chatni et al., 2009). When excited in the Soret band (395 nm) or either of the two Q bands (508 or 541 nm), the membrane displayed strong phosphorescence at 648 nm. The Soret band is within the blue region of the visible spectra,
while the Q bands are at the lower range of the green region. The highest intensity (measured as normalized relative fluorescence units, or NRFU) was 7.8 ± 0.1. This value was measured when the plate reader chamber was filled with nitrogen gas (≈ 0 kPa O2). As predicted by the Stern Volmer equation, the emission intensity was significantly lower when the chamber was filled with air (21 kPa O2) due to luminescent quenching by O2 (NRFU = 1.9 ± 0.2). This quenching effect is due to the excited triplet ground state of O2; excitation energy is released by a combination of non-radiative heat transfer, inter-particle collisions, and vibrational relaxation. Luminescent lifetime (i.e., phase shift) measured at a wavelength of 648 nm also showed O2 quenching (data not shown). For the CLPP studies below, excitation at the upper end of the Q band (≈ 545 nm) was selected based on the lower cost of filters and associated opto-electronic hardware. A number of O2 films were prepared with different concentrations of PtTFPP (while keeping polystyrene and chloroform concentrations constant). The average increase in emission intensity (NRFU) between 0 kPa and 21 kPa for all films was 70 ± 10% (Fig. 4A). Luminescent lifetime at an emission of 648 nm showed similar trends, but was more sensitive than intensity mode at low dye concentrations (Fig. 4B). The average increase in measured phase shift in 0 kPa and 21 kPa was 86 ± 7%. This significant increase in sensitivity (relative to NRFU) was due to the noise filtering capability of the phase-sensitive detection scheme in lifetime mode (Wolfbeis, 2004; McLamore et al., 2010; Chatni et al., 2009; Roche et al., 2010; Chaturvedi et al., 2013). During preliminary experiments, response time when changing from 0% to 21% O2 (or vice versa) was measured to be b60 s for all films. To maximize performance while limiting cost of the CLPP technique, 0.4% (w/w) membranes were used for all experiments to follow. For all CLPP analysis below, intensity mode was used to assess O2 dye films. As demonstrated in Fig. 4 and in the literature (Amao et al., 2000; Wolfbeis, 2004), lifetime mode analysis is more sensitive than intensity mode fluorescence. Additionally, lifetime mode analysis is
Fig. 3. A) Absorption spectra of PtTFPP dye immobilized in the bottom of a 96 well microtiter plate. B) Luminescence spectra of the sensing membrane after excitation at 545 nm. Solid line in panel B represents data collected in a nitrogen atmosphere (0 kPa O2); dashed line represents data collected in air (21 kPa O2). The emission intensity (measured in NRFU) is lower in the presence of O2 as described by the Stern Volmer principle.
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Fig. 4. A) Measured emission intensity (NRFU) at 645 nm for films exposed to 0 kPa (nitrogen gas) and 21 kPa. B) Measured phase shift (lifetime) at 0 kPa and 21 kPa O2. Lifetime mode measurements (panel B) are more sensitive than intensity mode measurements (panel A).
3.2. Comparison of PtTFPP to BDOBS plates In a preliminary study, CLPP using PtTFPP O2 plates prepared with a dye concentration of 0.3% (w/w) showed markedly lower levels of peak response compared to the commercial BDOBS O2 assay (see supplemental data, Fig. S1). Overall, peak response appeared to be limited to ~2 NRFU in the PfTFPP plates. To resolve this problem, PtTFPP plates were prepared with a dye concentration of 0.4% (w/w) to increase the maximum emission signal and the assays were repeated. When using PtTFPP plates with a dye concentration of 0.4%, there was no statistical difference in the maximum peak intensity (NRFU) when compared to BDOBS plates for the same soil slurries (p b 0.001; α = 0.05). For agricultural soils, supplementation of nitrogen or carbon increased average maximum intensity in all soils tested. Fig. 5 shows a representative transient plot showing peak NRFU response. Addition of supplemental nitrogen (10 mg L-1 (NH4)2SO4) caused a significant increase in peak aerobic respiration (23.2 ≈ 3.4%), but did not significantly change the time to peak (p b 0.001, α = 0.05). For all soils tested, addition of nitrogen alone caused an increase in peak NRFU of approximately 50%, while addition of casein, asparagine, and coumaric acid with nitrogen caused an increase in peak NRFU of 25–30% (Fig. 6). The largest increase was after the addition of nitrogen and mannose (72%). These trends are similar to those reported by Garland et al for similar soils using BDOBS plates (Garland et al., 2012), although the peak NRFU values in these samples are higher than those previously reported. For the Scrub and Hardwood soils, supplemental carbon addition did not always result in a significant increase in peak respiration. Mannose, coumaric acid and acetate significantly increased peak NRFU response, but casein and asparagine supplementation did not significantly increase peak NRFU. Regardless of N amendment, mannose, coumaric acid and acetate had the largest impact on SIR in Hardwood and Scrub soils. In Scrub soils, the major difference was a noted increase in asparagine metabolism when nitrogen was supplemented. While these minor differences in soil physiology are still being investigated, the important finding here is that no significant
differences were measured between the experimental PtTFPP and conventional BDOBS assays. The measured time associated with peak fluorescence (i.e., lag time) from samples treated with the PtTFPP dye method was not significantly different than with the BDOBS method (p b 0.0001, α = 0.05). Together with the peak response data, the new method appears comparable to the BDOBS method for CLPP (see supplemental data, Fig. S2 for time to peak data). The PtTFPP and commercial BDOBS plates were re-used seven times to estimate the shelf life and/or any sensor hysteresis. The average drift (i.e., percent change in output) was determined by calculating the mean percent change in peak response for each well in the assay, and then calculating the arithmetic mean of all peak NRFU responses (representing the net drift in sensor output). After each experiment, plates were rinsed, and soaked in DI water at 30 °C for 6 h, and air dried in the dark for 6 h. The threshold for performance was a 5% loss in performance (noted by a horizontal dashed line in Fig. 7). Based on this threshold, the PtTFPP plates may be re-used up to four times, while the BDOBs plates should not be used more than once. The reusability of BDOBS plates reported here was similar to results reported by Birmele et al (Birmele et al., 2006) characterizing planktonic Bacillus subtilis 1A2 and Pseudomonas aeruginosa PA01 respiration. This difference in performance between the two dye films is likely due to reduced photobleaching of platinum luminophores relative to
2.4 Scrub: Acetate (no N) Scrub: Acetate + N)
2.2 2.0
Emission [NRFU]
known to cause less photobleaching than intensity mode measurements. Photobleaching is a common problem with fluorescent dyes and causes signal drift. Although lifetime mode measurements are preferred if possible, intensity mode measurements were chosen for the following reasons: 1) a wealth of data exists in the literature for intensity mode measurements with BDOBS plates, 2) plate readers with lifetime mode capability are relatively new and therefore may limit the broad applicability of the technique, and 3) the lifetime of PtTFPP is different than the that of the ruthenium dye within the BDOBS plate, potentially imparting bias on the comparative study.
1.8 1.6 1.4 1.2 1.0 0.8 0
500
1000
1500
2000
2500
3000
Time [min] Fig. 5. Representative plot showing transient response of PtTFPP assay for an oak saw palmetto scrub soil sample with supplemented acetate. The closed circles show the transient response in the absence of supplemental nitrogen (10 mg L−1 (NH4)2SO4); the open triangles show the respiratory response with supplemental acetate and nitrogen.
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Percent change in peak response [%]
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PtTFPP plates BDOBS plates
0
-5
-10
-15
-20
-25 0
1
2
3
4
5
6
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8
Number of repeated uses [#] Fig. 7. Comparison of change in PtTFPP (0.4% w/w dye concentration) and BDOBS peak response during repeated use. The horizontal dotted line represents a 5% reduction in measured peak O2 response for all wells in the assay.
ruthenium-based dyes (Wolfbeis, 2004). In addition, the PtTFPP is nonionic and hydrophobic, reducing the leaching rate from the sensing membrane. Since output from all wells (both abiotic and biotic) were grouped together before calculating the percent change in output, the data represent the combined effects of photobleaching during measurement as well as hysteresis due to exposure of the dye to organic acids commonly found in soils (e.g., malate, citrate). If available, lifetime mode is recommended for all CLPP using the PtTFPP assay due to the enhance performance and increased longevity of the sensor. Available commercial systems such as the sensor dish reader (SDR) by Presens and MitoExpress© by LuxCel Biosciences. The SDR performs a similar oxygen analysis but is only available in 6 well or 24 well format. The SDR has the advantage of simultaneous analysis of pH and O2 within each, but has some disadvantages for soil analyses. In addition to prohibitive costs for soil analyses of large fields (where thousands of samples may be required), multi-parameter systems such as the SDR require complex three dimensional calibration curves and thus technician training. The MitoExpress© instrument also measures pH and O2, but is primarily intended for analysis of suspended cells and is not reliable for rapid multiplexing with natural soils. These challenges and others are discussed in detail in the review by Chaturvedi et al (Chaturvedi et al., 2013). Table 1 provides a basic cost summary for potential users of the technique. The total cost for preparing one 96 well plate for CLPP analysis in 2013, is approximately $15 USD (not accounting for potential reuse of the plate). Ordering supplies in bulk can reduce this cost by up to 8%. Use of platinum porphyrin dyes in opto-electronics is expected to increase in the coming decade, which will likely reduce the cost of assay preparation. The open source recipe described here provides soil scientists with a reliable recipe for preparing O2-sensitive gels used for physiological profiling of soil microbial communities.
Table 1 Cost summary in 2013 (in USD) for PtTFPP plates used for CLPP analysis. The total cost per plate is approximately $15 for the recipe with 0.4% dye.
Fig. 6. Comparison of PtTFPP (black bars) and BDOBS (gray bars) peak NRFU for samples in the presence and absence of various carbon/nitrogen substrates. O2-CLPP assays for (A) Agricultural soil, (B) Scrub soil, and (C) Hardwood soil. Carbon supplement contained 300 mg L−1 of indicated compound; (+N) indicates addition of 60 mg L1 (NH4)2SO4.
Item
Company
Pt(II) meso-tetra Pentafluorophenyl porphine BD Falcon 96 well plates Chloroform Polystyrene beads
Frontier Scientific PtT975 BD Biosciences Sigma Aldrich Sigma Aldrich
Product no.
353916 472476-1 L 39411-50G-F
Cost per plate [USD] $11.24 $3.02 $0.59 $0.12
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Validation of the new recipe presented here allows for continued development of the O2-based CLPP approach. As discussed in the introduction, this technique provides a less biased method for physiological profiling compared to previous formats (i.e., Biolog plates) due to lower substrate concentrations and elimination of redox dyes and proprietary growth supporting nutrients. However, the purpose of the CLPP approach, even with the O2-based platform, is rapid screening of the physiological capacity of microbial communities and not direct assessment of in situ physiological activity. Recent work with the BDoxy plates showed that the O2 peak responses between 6–24 h correspond to less than 1 to ~4 doublings (Garland et al., 2012). Therefore, the assay is useful for determining how communities are poised to utilize different types of substrate and what types of factors (e.g., N) may be limiting activity. This is particularly important when intensive spatiotemporal sampling and/or multiple experiment spatial treatments are of interest. When estimating specific rates of activity are of concern, alternative methods should be employed.
4. Conclusion Previous researchers have verified the utility of the O2-CLPP to assess the physiological status of microbial communities, particularly as applied to soil. To meet this need, a simple recipe was developed for fabrication of an open source O2-sensitive 96 well assay. The assay screens soil community physiology within a short period (≈24 h) by monitoring aerobic respiratory activity in microtiter wells using an oxygenquenched luminescent dye. Following the CLPP approach developed with commercial assays, substrate induced respiration and endogenous respiration were monitored for until a peak respiratory response is reached. The performance of the sensor technology is equivalent to a previously available commercial assay, but the sensor developed herein proved to be more robust and may be amenable to reuse.
Acknowledgments The authors would like to thank the Florida Agriculture Experimental Station (M.D. Dukes), the UF Excellence Award, and the IFAS Early Career Award (CRIS No. 005062) for funding. The authors also thank David Julian for the assistance with plate reader analysis.
Appendix A. Supplementary data Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.mimet.2013.10.016.
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