MIMET-04525; No of Pages 9 Journal of Microbiological Methods xxx (2014) xxx–xxx

Contents lists available at ScienceDirect

Journal of Microbiological Methods journal homepage: www.elsevier.com/locate/jmicmeth

3Q1

Ali Bozorg, Ian D. Gates, Arindom Sen ⁎

4

Department of Chemical and Petroleum Engineering, Schulich School of Engineering, University of Calgary, 2500 University Drive NW, Calgary, Alberta T2N 1N4, Canada

5 Q3 6 7 8 9 10

a r t i c l e

11 Q3 12 13 14 15 16

Keywords: Biofilm Porous medium Bioluminescence Biofilm saturation Relative hydraulic conductivity curve

O

R O

i n f o

F

2

Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties

a b s t r a c t

Article history: Received 9 August 2014 Received in revised form 24 November 2014 Accepted 26 November 2014 Available online xxxx

E

D

P

Biofilm formation in natural and engineered porous systems can significantly impact hydrodynamics by reducing porosity and permeability. To better understand and characterize how biofilms influence hydrodynamic properties in porous systems, the genetically engineered bioluminescent bacterial strain Pseudomonas fluorescens HK44 was used to quantify microbial population characteristics and biofilm properties in a translucent porous medium. Power law relationships were found to exist between bacterial bioluminescence and cell density, fraction of void space occupied by biofilm (i.e. biofilm saturation), and hydraulic conductivity. The simultaneous evaluation of biofilm saturation and porous medium hydraulic conductivity in real time using a non-destructive approach enabled the construction of relative hydraulic conductivity curves. Such information can facilitate simulation studies related to biological activity in porous structures, and support the development of new models to describe the dynamic behavior of biofilm and fluid flow in porous media. The bioluminescence based approach described here will allow for improved understanding and control of industrially relevant processes such as biofiltration and bioremediation. © 2014 Published by Elsevier B.V.

T

1

29

C

33 31 30 32

1. Introduction

35

Biofilms are structured communities of microorganisms embedded in a self-produced organic matrix of extracellular polymeric substances (EPS). In porous structures, which inherently have high surface area to volume ratios in comparison to non-porous structures, microbes can quickly colonize pore surfaces and form biofilms rather than remaining in a planktonic state (van Loosdrecht et al., 1990; Bouwer et al., 2000). With sufficient nutrient supply and metabolic waste removal, biofilms can progressively accumulate within a pore space making it increasingly difficult for fluids to flow through the porous structure (i.e. bioclogging) (Taylor and Jaffé, 1990; Cunningham et al., 1991; Thullner et al., 2002; Bozorg et al., 2011, 2012). In general, past studies have revealed that the permeability and porosity spatial distributions of a porous medium are related to the rate at which biofilms grow and spread within that medium. Biofilms are often considered to be undesirable as they can negatively impact industrial processes that rely on the flow of fluids in porous media. However, management of biofilm growth in porous media can improve the performance of those industrial and environmental processes that rely on biofilms to achieve a process goal, such as in situ bioremediation (Thomas and Ward, 1989; Madsen, 1991; Singh et al., 2006), biobarrier containment (Kim et al., 2006), wastewater treatment (Nicolella et al., 2000), enhanced oil recovery (Lappin-Scott

42 43 44 45 46 47 48 49 50 51 52 53 54 55 56

R

R

40 41

N C O

38 39

U

36 37

E

34

⁎ Corresponding author. Tel.: +1 403 210 9452; fax: +1 403 284 4852. E-mail address: [email protected] (A. Sen).

et al., 1988), and carbon sequestration (Mitchell et al., 2009). Development of robust methods to engineer biofilms in porous structures requires comprehensive knowledge of the processes that affect their spatiotemporal development under different flow conditions. However, due to an inadequate understanding of the interactions between porosity and permeability, biofilm growth kinetics, multiphase flow effects, spatial variations of cell nutrients, and impact of medium heterogeneity, field scale applications of biofilm based processes are still unpredictable (Gerlach and Cunningham, 2010). Different models have been used to simulate biofilm growth in porous media (Baveye and Valocchi, 1989; Vandevivere et al., 1995; Clement et al., 1996; Thullner et al., 2004; Kim and Whittle, 2006; Bozorg et al., 2011). The general approach used in these models has been to incorporate relationships that link porosity to hydraulic conductivity by treating biofilm as an emerging solid phase that changes the intrinsic porosity and permeability of a medium. Whereas such models can qualitatively reproduce experimental results, they are limited due to the lack of reliable correlations between porosity and permeability (Baveye and Valocchi, 1989; Clement et al., 1996; Thullner et al., 2002; Bozorg et al., 2011). Recently, Bozorg et al. (2011) introduced a new macroscopic approach to model biofilm spatiotemporal development in porous media by treating biofilm as a high viscosity liquid phase that shares pore space with a low viscosity aqueous phase. In that study, efforts were made to quantify effective conductivities of water and biofilm phases via relative permeability curves based on biofilm saturation (i.e. fracture of pore space occupied by biofilm). However, calibration of the parameters used in this approach is challenging since it

http://dx.doi.org/10.1016/j.mimet.2014.11.015 0167-7012/© 2014 Published by Elsevier B.V.

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

17 18 19 20 21 22 23 24 25 26 27 28

57 58 59 60 61 62 63 64 65 66 67 68 69 70 71 72 73 74 75 76 Q4 77 78 79 80 81 82 83

The bioluminescent bacterial strain, porous medium, imaging system, and fluid application system have been described in detail elsewhere (Bozorg et al., 2012), and so will only be described here briefly.

120

2.1. Bacterial strain and culturing conditions

121 122

This study used the bioluminescent reporter strain Pseudomonas fluorescens HK44 (hereafter referred to as HK44) obtained from the University of Tennessee Center for Environmental Technology (University of Tennessee, Knoxville, TN). The strain harbor had been genetically modified previously by transposon insertion of the salicylate-inducible luxCDABE gene cassette and a tetracycline resistance marker, thereby enabling it to produce luciferase when salicylate is present, which in turn causes the cells to luminesce (King et al., 1990). HK44 was grown for 18 h on an orbital shaker (Heidolph Unimax 2010, Germany) at 150 rpm and 25 °C in an oxygen-saturated, nitrate-free growth medium (pH = 7.20 ± 0.05) consisting of MgSO4, 0.4 g/L; CaCl2·2H2O, 0.1 g/L; NH4Cl, 0.4 g/L, NaCl, 8 g/L; KCl, 0.2 g/L; NaH2PO4, 1.15 g/L; K2HPO4, 0.26 g/L; HCl, 0.00366 g/L; FeSO4·7H2O, 0.021 g/L; H3BO3, 0.0003 g/L; MnCl2·4H2O, 0.001 g/L; CoCl2·6H2O, 0.0019 g/L; NiCl2·6H2O, 0.00024 g/L; CuCl2·2H2O, 0.00002 g/L; Na2EDTA·2H2O, 0.01 g/L; ZnSO4·7H2O, 0.00144 g/L; and Na2MoO4·2H2O, 0.00036 g/L. Glucose was added to a final concentration of 1.0 g/L as the main carbon source. After being examined for bacterial bioluminescence, actively growing cultures were concentrated by centrifugation (Beckman Coulter®, X-22R), suspended in a glycerol stock culture, and stored at − 80 °C in 1 mL aliquots. All experiments were inoculated directly using cells from the frozen stock. To induce luminescence, an induction medium was prepared by removing all phosphate sources from the growth medium, and adding 0.1 g/L (final concentration) of salicylate (Bozorg et al., 2012). Also, all

109 110 111 112 113 114

123 124 125 126 127 128 129 Q2 130 131 132 133 134 135 136 137 138 139 140 141 142 143 144 145

C

107 108

E

105 106

R

103 104 Q6

R

101 102

O

99 100 Q5

C

97 98

N

95 96

U

93 94

In this study, translucent acid-washed glass beads with diameters ranging between 425 and 600 μm (Sigma-Aldrich, G8772) were used as porous medium. The porosity of the bead pack was determined gravimetrically using the mass of water within the pores (Bozorg et al., 2012). Prior to use, the glass beads were washed twice with distilled water to remove fines, and autoclaved at 121 °C for 20 min.

149 150

2.3. Dissolved oxygen measurement

155

F

118 119

91 92

148

An oxygen microelectrode (MI-730, Microelectrodes Inc.) was used to monitor oxygen concentration. In the flow chamber, an inline microelectrode was placed at the chamber outlet for real-time evaluation of oxygen concentration during biofilm growth and bacterial bioluminescence under flow conditions. The electrode was connected via an amplifier to an eDAQ Data Acquisition System (eDAQ PTY LTD, Australia) interfaced to a computer through a USB-port. Two separate media with 0% and 100% oxygen saturations, respectively, were used to calibrate the electrode.

O

117

90

2.2. Porous media

R O

2. Materials and methods

88 89

2.4. Imaging

P

116

86 87

media used in this study were supplemented with 30 mg/L tetracycline 146 (EMD Chemicals, OmniPur® EM-8990) to ensure plasmid maintenance. 147

151 152 153 154

156 157 158 159 160 161 162 163 164 165

A 14-bit digital charged-coupled device (CCD) camera (Progres MFcool, Jenoptik, Germany) equipped with a Computar Megapixel lens with 35 mm focal length and f/1.4 focal ratio was used to capture grayscale bioluminescence images. The CCD camera was positioned 40 cm above the media. All bioluminescence experiments were conducted with the equipment in a dark box to eliminate the effects of extraneous light. Throughout the experiments, all images were acquired with a 5 minute exposure time with a fully open aperture. The imaging process, and method to evaluate BI were previously validated and are described in detail in Bozorg et al. (2012).

166

2.5. Experimental procedures

176

To accomplish the goals of this project using imaging technology, it was necessary to correlate bacterial bioluminescence to bacterial cell density and biofilm saturation. As bioluminescence has been reported to be a function of oxygen concentration, we evaluated the impact of oxygen on detected bioluminescence in our samples under induction conditions and used this information to ensure that O2 was not limiting our observations. We then determined the impact of the packing material on the observed bioluminescence from both planktonic bacteria as well as bacteria within biofilms, and subsequently used this knowledge to study how biofilms affect fluid flow in porous media. We finally used established correlations to nondestructively evaluate biofilm saturation under flow conditions. The methods used to carry out these studies are described here.

177

2.5.1. Effect of oxygen on bacterial bioluminescence To evaluate the effect of oxygen concentration on the bioluminescence intensity of HK44, liquid batch cultures of these cells were grown on a rotary shaker (Heidolph Unimax 2010, Germany) at 150 rpm and room temperature. After 18 h, bacteria were harvested by centrifugation at 5000 relative centrifugal force (rcf) for 20 min and washed twice with phosphate buffered saline (PBS). Known quantities of the harvested cells were then redistributed in batch cultures containing induction medium, and subsequently, the oxygen concentration and bioluminescence intensity of induced cells were monitored. All cell densities were evaluated by measuring light absorbance at 550 nm in a spectrophotometer (DU 730, UV/Vis Spectrophotometer, Beckman

190

T

115

requires simultaneous evaluation of biofilm evolution and hydraulic conductivity. A majority of studies on biofilm growth in porous media involve destructive sampling, i.e. removing biofilm from an experimental apparatus for characterization purposes (Taylor and Jaffé, 1990; Cunningham et al., 1991; Seki et al., 2006). Evaluation typically occurs only at the end of an experiment, which is not conducive to studying how biofilms evolve and affect fluid flow in real time. Real-time quantification of biofilm characteristics and porosity and permeability have been a subject of interest in porous media based processes for many years (Oostrom et al., 1998; Niemet and Selker, 2001; Seymour et al., 2004; Bozorg et al., 2012). Recently, new approaches have been developed that use bioluminescence emitted by certain natural and engineered microorganisms to monitor microbial processes in natural and engineered environments (Burlage et al., 1990; Shaw et al., 1992; Ripp et al., 2000; Uesugi et al., 2001; Sharp et al., 2005; Bozorg et al., 2012). For instance, Sharp et al. (2005) used naturally luminescent bacteria in a flat-plate flow chamber to study biofilm growth under flow; whereas bacterial bioluminescence was used to track biofilm development, no quantification was made of the detected bioluminescence intensity (BI). In another experimental study, Bozorg et al. (2012) used a CCD camera to monitor growth of biofilm forming bioluminescent bacteria in a translucent porous medium. They demonstrated that it may be possible to use inducible bacterial bioluminescence to nondestructively evaluate cell density and hydraulic properties in porous media. The objective of the research that will be described here was to develop an approach to quantify, nondestructively and in real-time, porous medium hydraulic conductivity and biofilm saturation, and to then use these parameters to develop relative hydraulic conductivity curves for the flowing aqueous phase. The ability to determine such information will enable better understanding of interactions between biofilm growth and fluid flow in porous structures.

D

84 85

A. Bozorg et al. / Journal of Microbiological Methods xxx (2014) xxx–xxx

E

2

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

167 168 169 170 171 172 173 174 175

178 179 180 181 182 183 184 185 186 187 188 189

191 192 193 Q7 194 195 196 197 198 199 200 201

A. Bozorg et al. / Journal of Microbiological Methods xxx (2014) xxx–xxx

221 222 223 224 225 226 227 228 229 230 231 232 233 234 235 236 237 238 239 240

2.5.3. Bioluminescence of biofilm bacteria in porous medium under static conditions To determine the bioluminescence activity of biofilm bacteria in porous medium under static conditions, 33 PPs were inoculated through immersion in a cell suspension (6 × 105 cells/mL) for 8 h to provide sufficient time for bacterial attachment to the porous matrix. Subsequently, each PP was removed from the cell suspension and placed in a Petri dish containing 28 mL of sterile growth medium to allow for bacterial proliferation and biofilm formation within the porous medium. Thereafter, every 12 h over the course of 8 days, three PPs were transferred to a second Petri dish containing fresh induction medium, where the bioluminescence of the bacteria in the biofilm within the porous medium was measured. The biofilm from each PP was then harvested and the bacterial cell density determined (see Section 2.7 for details). The obtained data were used to develop a correlation between measured bioluminescence and the number of cells in a biofilm within a translucent porous medium. Note that it has been previously reported that HK44 cells retain their viability under static conditions over a period of 8 days (Bozorg et al., 2012). In addition, in this study, daily samples obtained from the biofilm cells revealed similar bacterial proliferation kinetics for not less than 8 days.

F

O

K¼

a)

qL Ah

R O

220

243 244 245 246 247 248 249 250 251 252 253 254 255 256 257 258 259 260 261

ð1Þ

P

219

241 242

where q is volumetric flow rate of the medium, A is cross-sectional area 263 of the porous medium, and h is hydraulic head difference applied over a length L. During the induction phase, growth medium was replaced by 264 induction medium to induce the lux-genes. 265

D

217 218

E

215 216

T

213 214

C

211 212

E

209 210

R

207 208

R

206

2.5.2. Bioluminescence of planktonic bacteria in porous medium A total of 72 Porous Packs (PPs), each consisting of a 1 cm thick cell strainer (BD Falcon™ 352360) with 100 μm nylon mesh, filled with dried glass beads (Fig. 1a) were used to determine the impact of the porous medium packing material on the measurement of bioluminescence emitted by planktonic cells. Each PP was placed in a separate 60 cm Petri dish filled with induction medium containing a known density of planktonic HK44 cells (determined spectrophotometrically). After a sufficient period of time for the planktonic cells to fully penetrate the porous medium, a bioluminescence image was recorded of a given PP in its Petri dish (Fig. 1a). An image was also recorded of the liquid culture in the same dish in a region away from the PP. The difference in the recorded BI between the PP and its surrounding liquid culture in the dish was used to evaluate the effect of the porous medium material on detected BIs. Obtained results were also used to correlate BIs with cell density in both porous medium and liquid culture.

N C O

204 205

2.5.4. Bioluminescence of biofilm bacteria in porous medium under flow conditions To simultaneously evaluate biofilm bioluminescence and its effects on hydraulic conductivity, bacterial cells were allowed to grow and form biofilm under flow conditions in a packed tube (PT) device (Fig. 1b). Each PT consisted of a rectangular clear acrylic tube (0.95 cm internal width) filled to a height of 2 cm with dried glass beads which were held in place by 0.1 mm nylon mesh screens at the inlet and outlet. Each PT was filled with sterile growth medium via a peristaltic pump (Gilson Minipuls 3) while orientated in a vertical position. The same pump was then used to add 0.6 mL of cell culture inoculum (6× 105 cells/mL) to each PT. We previously verified that the pump did not adversely impact bacterial cell viability. To minimize the possibility of washout, bacteria were allowed to attach to the glass beads overnight at room temperature. Continuous flow of growth medium was then initiated by using a Mariotte tube (SMS®, Arizona, USA) to maintain a constant pressure head. At regular intervals, each PT was simultaneously evaluated for microbial bioluminescence and fluid flow rate. The flow rate was recorded using a calibrated in-line flowmeter (BEL-ART Riteflow®) at the outlet of the chamber. The hydraulic conductivity (K) of the porous medium was determined by:

Coulter®) which had been calibrated previously by using a cell counting chamber (Hemacytometer Set, Hausser Scientific).

2.6. Determining biofilm saturation

266

To calculate biofilm saturation (i.e. fraction of original void space occupied by biofilm in a sample of porous medium), the volume of biofilm (Vb) in that sample is required. However, as direct measurement of biofilm volume in porous samples is not feasible, it was instead estimated by using biofilm dry density (ρbd), defined as the mass of dry biofilm per unit volume of hydrated biofilm (including water), and the mass of dried biofilm (mdb) after removing all water content as per the equation:

267

Vb ¼

mdb : ρbd

268 269 270 271 272 273 274

ð2Þ 276

b)

U

202 203

3

Fig. 1. (a) Porous Packs (PPs) were used to evaluate bacterial bioluminescence in porous media and liquid culture. Each PP consisted of a 100 μm cell strainer, filled with glass beads, and was placed in a Petri dish containing growth or induction medium with HK44 cells. (b) Packed tubes (PT), each consisting of a rectangular clear acrylic tube (0.95 cm internal width) filled to a height of 2 cm with dried glass beads. PTs were used to grow biofilm under a constant pressure gradient in order to study hydraulic conductivity variations.

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

A. Bozorg et al. / Journal of Microbiological Methods xxx (2014) xxx–xxx

7.843 × 10−15 g per cell based on the PUTK21 plasmid used in HK44 and DNA of the natural bacteria (King et al., 1990). A standard curve, correlating mass of DNA to bacterial cell population size, was generated by manual cell counting in a counting chamber (Hemacytometer Set, Hausser Scientific). Cell numbers (at least 360 bacteria per slide for statistical relevance) were counted in triplicate for each sample.

316 317

3. Results and discussion

322

3.1. Oxygen effects

323

Lux gene dependent bioluminescence has been shown to be an aerobic process (Meighen and Dunlap, 1993; Uesugi et al., 2001). Hypoxia, therefore, leads to a decrease in bioluminescence. The kinetic values for these processes have been studied extensively for both immobilized (Webb et al., 1997) and suspended (Kelly et al., 2003) cells. Thus, prior to investigating any application of lux gene dependent bioluminescence, it was deemed important to determine the oxygen levels needed to make sure it did not limit bacterial bioluminescence during subsequent experiments. To accomplish this, planktonic cultures of HK44 were inoculated into induction medium (1.7 × 108 cells/mL) which had been saturated with air using an aquarium air pump (Whisper® 196 Tetra Holding, US). The vessel was operated in batch mode, and the oxygen concentration in the culture medium and bioluminescence were simultaneously measured over time. As shown in Fig. 3a, bioluminescence was not detected for approximately the first 25 min. This delay may be attributed to the time required to activate the cellular salicylate transport mechanism (King et al., 1990), as well as the time required to accumulate a threshold quantity of luciferase needed for bioluminescence detection (Kelly et al., 2003). During this period, there was not much change in oxygen concentration indicating that the cellular oxygen uptake rates were minimal. However, after 25 min, the oxygen uptake rate increased sharply, coinciding with a rapid rise in BI. These observations were consistent with other experimental studies (Uesugi et al., 2001). Also, as expected, our results revealed that low oxygen concentrations (present after approximately 90 min) negatively affect bioluminescence levels (Fig. 3a). Similar trends were observed for other cell densities ranging from 1 × 105 to 2 × 108 cells/mL (i.e. BIs increased up to a maximum value, remained constant for a period of time, and then decreased after the dissolved oxygen concentration dropped below a critical level). As shown in Fig. 3a, stable oxygen consumption rates during maximum bacterial bioluminescence, and the corresponding critical oxygen concentration (e.g. 3.6 μmol/min and 50 μmol oxygen, respectively) imply that higher oxygen concentrations could potentially increase the bioluminescence period. As shown in Fig. 3b, further investigation revealed that saturating the medium with pure oxygen (as opposed to air that was used for the results shown in Fig. 3a) significantly increased the constant bioluminescence period by up to 200 min while keeping the maximum bioluminescence value intact. Therefore, to reduce the negative impact that oxygen availability had on bacterial bioluminescence, all media were saturated with pure oxygen (Praxair Inc.) for all remaining experiments.

324 325

3.2. Cell density evaluation

366

One goal of this work was to determine cell density within a biofilm by simply measuring bioluminescence intensity. Therefore, it was important to simultaneously quantify these two parameters and determine a correlation between them. Fig. 4 shows that lux gene dependent bioluminescence can be correlated to bacterial cell densities, whether the cells are (i) in planktonic form in liquid culture, (ii) in planktonic form within a translucent porous medium, or (iii) within a biofilm in a translucent porous medium. For the planktonic cells, a total of 72 samples with different cell densities were studied to obtain a relationship between bioluminescence activity of planktonic cells

367

308

At the end of an experiment, biofilm covered porous media, from either PPs or PTs, were placed into separate test tubes, each containing a known volume of PBS, and vortexed and sonicated repeatedly to detach biofilm from the glass beads (Heersink, 2003). These samples were used to determine the bacterial cell numbers by measuring the total DNA weight (Qubit® dsDNA Assay Kit, Invitrogen, Oregon, USA). Since a single cell strain was used, the total DNA weight could be correlated to bacterial concentration. Total mass of HK44 DNA was calculated to be

294 295 296 297 298 299 300 301 302 303 304 305

309 310 311 312 313 314 315

C

E

R

292 293

R

291

O

289 290

C

287 288

N

285 286

U

283 284

P

2.7. Bacterial cell enumeration

281 282

D

307

279 280

T

306

For gravimetric measurements, bacterial cells were placed in growth medium (3 replicates) and allowed to form biofilm on glass slides (Fig. 2). After 12 days, the biofilm thickness (Lb), was measured with an optical microscope (Axio Observer.A1, Zeiss, Germany) according to the method of Bakke and Olsson (1986). Nine optical-thickness measurements were taken at regular intervals for each slide and then averaged to obtain a mean biofilm thickness. Also, surface coverage of a slide was estimated by image analysis from digital images taken using the previously described imaging system. Using MATLAB® Image Processing Toolbox (MATLAB® R2009a, MathWorks Inc.), a threshold value (Otsu, 1979) was applied to biofilm images so that those areas covered with biofilm appeared white, while the remaining area appeared black. The relative surface coverage of the biofilm was then calculated as the proportion of white to the total area. This thresholding approach has been commonly used with images of biofilm to discern the biological component from the background (Stoodley et al., 1998; Bozorg et al., 2012). Subsequently, each slide was dried at 85 °C for 24 h, and after being weighed, the mass of the clean slide (taken before the experiment) was deducted to calculate the biofilm dry mass (mdb). The measured volume and mass values were then used as per Eq. (2) to calculate the biofilm dry density. Since identical growth conditions were used throughout the experiments to grow biofilm, a consistent dry density was assumed in all conditions for the biofilm formed in the porous media (Characklis and Cooksey, 1983). Therefore, to estimate the biofilm volume, each PP was weighed prior to inoculation. After inoculation and at different time intervals, inoculated PPs were removed from the Petri dishes, dried at 85 °C for 24 h and weighed to determine dry mass of biofilm. By using the measured biofilm dry mass and biofilm dry density, the fraction of pore space occupied by biofilm (i.e. biofilm saturation) was calculated.

277 278

E

Fig. 2. Biofilm coated glass slides used to determine volume of biofilm. Bacterial cells were allowed to attach and form biofilm for 12 days on the glass slide and subsequently, biofilm thickness and surface area of the covered glass slide was measured.

R O

O

F

4

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

318 319 320 321

326 327 328 329 330 331 332 333 334 335 336 337 338 339 340 341 342 343 344 345 346 347 348 349 350 351 352 353 354 355 356 357 358 359 360 361 362 363 364 365

368 369 370 371 372 373 374 375 376

5

R

E

C

T

E

D

P

R O

O

F

A. Bozorg et al. / Journal of Microbiological Methods xxx (2014) xxx–xxx

378 379 380 381 382 383 384 385 386 387 388 389 390 391 392 393 394 395 396 397

either in liquid culture or in porous medium. These experiments were carried out by placing a PP in a Petri dish containing medium inoculated with a known quantity of cells (i.e. a known liquid cell density). As shown in Fig. 4a, bioluminescence was measured in those parts of the dish where the PP was present (porous media) as well as those parts of the plate where the PP was not present (liquid culture). In this way, the impact of the porous medium on the measured bioluminescence of planktonic cells could be discerned. Fig. 4c shows results from a number of these samples, representing the entire range of planktonic cell numbers tested (results from all 72 samples were not included in this figure to maintain clarity). Not surprisingly, the bioluminescence detected for planktonic cells within the porous medium was lower when compared to cells in liquid culture. This can be attributed to the lower number of cells per unit volume of porous medium compared to liquid culture, as well as shielding of emitted bioluminescence by the packing material. To evaluate the shielding effect of the packing material, the porosity of the medium (calculated gravimetrically to be 38%) was used to first determine the actual number of cells within PPs. Based on these values, the expected BIs in the absence of any shielding by the porous medium were then calculated (using the correlation obtained in Fig. 4c for BI of

U

377

N C O

R

Fig. 3. (a) Effect of oxygen saturation on BI of planktonic HK44 cells (1.7 × 108 cells/mL) in induction medium saturated with sterile air. (b) Bioluminescence response of planktonic HK44 cultures with different population densities in induction medium which had been fully saturated with pure oxygen. Both experiments were conducted in batch mode, with oxygen concentrations and bioluminescence levels being measured simultaneously over time.

planktonic bacteria in liquid culture) and plotted against detected BIs (Fig. 5a). As anticipated, a linear relationship was observed between detected BIs and the calculated expected values with the proportionality constant of 0.63. This implies that, on average, the packing material shielded 37% of the emitted bacterial bioluminescence in these experiments. These data were used to calculate a shielding coefficient (also shown in Fig. 5a) using Eq. (3): Shielding coefficient Calculated BI without shielding −Detected BI : ¼ Calculated BI without shielding

398 399 400 401 402 403 404 405

ð3Þ 407

As shown in Fig. 5a, it is apparent that the degree of shielding by the packing material was more significant at lower BI values, and decreased from a maximum value of 0.85 to 0.3 when the calculated BI increased from 1.6 to 19.2. The combined impact of cells being in a biofilm and additionally within packing material on the measured bioluminescence is also shown in Fig. 4b and c. It is apparent that for a given number of cells, the measured bioluminescence from a biofilm within a porous medium

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

408 409 410 411 412 413 414

A. Bozorg et al. / Journal of Microbiological Methods xxx (2014) xxx–xxx

D

P

R O

O

F

6

424 425 426 427 428 429 430 431 432 433 434 435 436 437 438 439 440 441 442 443 444

E

R

R

422 423

O

420 421

C

418 419

was lower than from planktonic cells either within a PP or liquid culture. This result was expected; in biofilms, microorganisms are enclosed in a self-produced EPS that contributes to the overall shielding effect. To evaluate this biofilm shielding effect, the detected BI values of biofilm cells in the porous medium were plotted against the BI of planktonic cells in the same porous medium calculated as though there was no biofilm shielding (using the expression determined for this condition in Fig. 4c). As shown in Fig.5b, a linear correlation was again observed between the detected BI and that calculated without the effect of biofilm shielding. The proportionality constant of 0.42 indicates that on average, 58% of the emitted bacterial bioluminescence was shielded from detection due to being within a biofilm. The lower proportionality constant in Fig. 5b compared to 5a indicates that biofilm shielding had more impact on detected bioluminescence than the packing material used in these experiments. Fig. 5b also shows a calculated biofilm shielding coefficient (calculated using Eq. (3)) as a function of the calculated BI that would be expected if biofilm shielding was negligible. The obtained profile indicates that during the initial phase of biofilm formation in porous media, when the number of cells (and thus, measured BIs) are low, the shielding coefficient increased and reached a maximum value of 0.64 at a calculated BI of 6.1 (which corresponded to a detected BI of 2.3 and 5.6E + 06 cells/mL). The coefficient then remained constant until the calculated BI increased to 10.78 (which corresponded to a detected BI of 4.6 and 3.5E + 07 cells/mL), and then gradually declined to 0.5. This profile shows the impact of EPS on detected BI at different biofilm growth stages. As at the beginning, when the amount of EPS compared to the number of cells in a porous medium is low, its impact on emitted bacterial bioluminescence would also be low. However, as the biofilm grows and EPS occupies more pore spaces, the detected bioluminescence is

N

416 417

U

415

C

T

E

Fig. 4. (a) A typical image of a porous pack in a Petri dish containing medium inoculated with bioluminescent cells. The liquid culture refers to the lighter region in the well, outside the PP. The darker area (outlined with a dashed black circle) refers to the porous medium. (b) A typical image of a PP containing a biofilm (outlined here by a dashed white perimeter) all within a Petri dish. Note that the liquid medium in the Petri dish was not inoculated with cells. (c) Measured bioluminescence intensity of planktonic cells in liquid culture alone (single reading) (▲), planktonic cells in liquid culture within a translucent porous medium (single reading) (♦), and cells in a biofilm formed within a translucent porous medium (average of 3 replicates) (●). For clarity and to compare the BIs, bioluminescence images of 3 encircled points corresponding to the planktonic cells in liquid culture and porous medium (8.6 × 107 cells/mL) and biofilm cells in porous medium (8.9 × 107 cells/mL) are illustrated.

more adversely affected (Fig. 4b). In addition, when compared with the packing material related shielding coefficient, the higher shielding effect of EPS may be due in part to limited oxygen diffusion in dense biofilm regions with high levels of EPS. Whereas packing material and biofilm affect the detected BI values, Fig. 4c reveals a power relationship between cell density and detected BI for all three sets of data. Such nonlinear relationships between metabolic activity (e.g. bacterial bioluminescence) and number of cells have been observed in many biological processes, such as metabolic rates of entire organisms, maximal population growth rate, concentration of metabolic enzymes, and sizes of biological structures, with allometric scaling exponents ranging from 1.3 to 4 (West and Brown, 2005). Regression of the data showed that all the power functions possessed a similar exponent value of approximately 0.38, with different scaling factors, implying that bioluminescence exhibits the same behavior with respect to cell populations in all three sets of data while the scaling factors indicate environmental effects, including packing material and EPS shielding, on detected BIs.

445

3.3. Biofilm saturation

463

To correlate the impact of biofilm growth on fluid flow through a porous medium under different conditions, it was first necessary to find a way to measure biofilm saturation. Based on the measured biofilm thickness (Fig. 2), area of covered surface, and dry mass of biofilm, the biofilm dry density was calculated to be equals to 42.1 (± 6%) mg/mL which is consistent with values of 30 to 80 mg/mL reported in the literature (Beyenal et al., 1998; Garcia Lopez et al., 2003). The mass of dry biofilm was measured by finding the difference between the dry mass of the PP prior to inoculation and after the formation of biofilm. The mass of the

464 465

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

446 447 448 449 450 451 452 453 454 455 456 457 458 459 460 461 462

466 467 468 469 470 471 472

7

T

E

D

P

R O

O

F

A. Bozorg et al. / Journal of Microbiological Methods xxx (2014) xxx–xxx

479

R

R

477 478

N C O

475 476

dry biofilm together with the dry density was used to calculate the biofilm volume in the porous medium, which in turn allowed for biofilm saturation to be determined. Since the BI of each PP was recorded prior to drying, it was possible to develop a correlation between biofilm saturation and BI. As shown in Fig. 6, these two parameters could be correlated using a power law relationship. Under similar growth and induction conditions, this power law was used in conjunction with bioluminescence

measurements to estimate biofilm saturation in porous medium in the re- 480 maining experiments. 481 3.4. Hydraulic conductivity

482

An increase in biofilm saturation reduces the void space available 483 for fluid flow, and thus, lowers hydraulic conductivity. In the present 484

U

473 474

E

C

Fig. 5. (a) The detected BI of planktonic cells in porous medium as a function of the BI that would have been expected had the porous medium not provided shielding. Also shown is an overall shielding coefficient as calculated using Eq. (3). (b) The detected BI of biofilm cells within a porous medium shown as a function of the BI that would have been expected had the biofilm itself not provided shielding. Also shown is an overall shielding coefficient.

Fig. 6. Seven bioluminescence images captured from PPs are illustrated (scale indicates increasing BI from 0 to 12). Using biofilm dry density, biofilm saturation was evaluated gravimetrically in each PP. The results revealed that a power function provides a good fit between calculated biofilm saturation and recorded BI.

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

A. Bozorg et al. / Journal of Microbiological Methods xxx (2014) xxx–xxx

R

E

C

T

E

D

P

R O

O

F

8

489 490 491 492 493 494 495 496 497 498 499 500 501 502 503 504 505

C

N

487 488

study, significant reductions in hydraulic conductivities were observed during biofilm growth in PTs (Fig. 7a), consistent with the literature (Cunningham et al., 1991; Lappin-Scott et al., 1988; Seki et al., 2006; Bozorg et al., 2012). The correlation observed between biofilm saturation and BI (Fig. 6) suggested that bacterial bioluminescence may also be used for real-time evaluation of porous medium hydraulic conductivity. To investigate such a relationship, Eq. (1) was used to determine hydraulic conductivity of porous medium based on the applied pressure head and the fluid flux through the PTs. Immediately after the flux determination, a bioluminescence image was recorded from the induced cells and subsequently, relative hydraulic conductivities (ratio of effective hydraulic conductivity to the saturated hydraulic conductivity of the clean porous medium) were plotted as a function of evaluated BIs (Fig. 7a). The exponential relation observed between these two parameters indicates that measured bacterial bioluminescence can be used to estimate the hydraulic conductivity in real time. Hydraulic conductivity has previously been reported to be exponentially related to porosity (Clement et al., 1996). Assuming that the total porosity (φ) consists of a void porosity (εv) and an occupied biofilm fraction, several authors (Ives and Pienvichitr, 1965; Okubo and

U

485 486

O

R

Fig. 7. (a) Relative hydraulic conductivity of porous media versus BI of each packed tube. (b) Relative hydraulic conductivity of packed tubes plotted against evolving biofilm saturation (assessed by using bacterial bioluminescence) under flow conditions. The results indicate that the published Clement model (dashed line) tends to overestimate hydraulic conductivities at similar biofilm saturation levels. However, an expression equivalent to the Clement model but with a higher exponent (n = 6.33) provides a good fit.

Matsumoto, 1979; Knapp et al., 1988; Clement et al., 1996) proposed 506 the following relationship: 507 K K initial

¼

 n εv ϕ

nN0

ð4Þ

where Kinitial is initial (clean) hydraulic conductivity and n is an exponent that varies between different models based on the microgeometrical properties of the porous medium and morphology of the biofilm. By assuming homogenous and uniform biofilm growth on solid particles, Clement et al. (1996) developed a relationship identical to Eq. (4) and found that for typical sandy materials, n had a value of 19/6 (~3.167). By using the correlation between BI and biofilm saturation previously determined in PPs, relative hydraulic conductivities could now be obtained as a function of biofilm saturation (Fig. 7b). The results revealed that at biofilm saturation levels of less than about 4%, there is negligible impact on hydraulic conductivity. However, N 4%, the hydraulic conductivity dropped with increasing saturation as expected. Experimental results were also compared with the Clement model to see whether estimated biofilm saturation and measured hydraulic

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

509 510 511 512 513 514 515 516 517 518 519 520 521 522

A. Bozorg et al. / Journal of Microbiological Methods xxx (2014) xxx–xxx

Acknowledgment

557 559

The authors acknowledge financial support from the S. M. Blair Family Foundation, Natural Sciences and Engineering Research Council of Canada (NSERC), and Alberta Innovates — Technology Futures.

560

References

561 562 563 564 565 566 567 568 569 570 571 572 573 574 575 576 577 578 579 580 581 582 583 584 585 586 587 588 589

Bakke, R., Olsson, P.Q., 1986. Biofilm thickness measurements by light microscopy. J. Microbiol. Meth. 5, 93–98. Baveye, P., Valocchi, A., 1989. An evaluation of mathematical models of the transport of biologically reacting solutes in saturated soils and aquifers. Water Resour. Res. 25, 1413–1421. Beyenal, H., Tanyolac, A., Lewandowski, Z., 1998. Measurement of local effective diffusivity in heterogeneous biofilms. Water Sci. Technol. 38, 171–178. Bouwer, E.J., Rijnaarts, H.H.M., Cunningham, A.B., Gerlach, R., 2000. Biofilms in porous media. In: Bryers, J.D. (Ed.), Biofilms II: Process Analysis and Applications. WileyLiss, New York, USA, pp. 123–158. Bozorg, A., Sen, A., Gates, I.D., 2011. A new approach to model the spatiotemporal development of biofilm phase in porous media. Environ. Microbiol. 13, 3010–3023. Bozorg, A., Gates, I.D., Sen, A., 2012. Real time monitoring of biofilm development under flow conditions in porous media. Biofouling 28, 937–951. Burlage, R.S., Sayler, G.S., Larimer, F., 1990. Monitoring of naphthalene catabolism by bioluminescence with nah-lux transcriptional fusions. J. Bacteriol. 172, 4749–4757. Characklis, W.G., Cooksey, K.E., 1983. Biofilms and microbial fouling. Adv. Appl. Microbiol. 29, 93–127. Clement, T.P., Hooker, B.S., Skeen, R.S., 1996. Macroscopic models for predicting changes in saturated porous media properties caused by microbial growth. Ground Water 34, 934–942. Cunningham, A.B., Charaklis, W.G., Abedeen, F., Crawford, D., 1991. Influence of biofilm accumulation on porous media hydrodynamics. Environ. Sci. Technol. 25, 1305–1311. Garcia Lopez, L.A., Veiga, M.C., Nogueira, R., Aparicio, A., Melo, L.F., 2003. A technique using a membrane flow cell to determine average mass transfer coefficients and tortuosity factors in biofilms. Water Sci. Technol. 47, 61–67. Gerlach, R., Cunningham, A.B., 2010. Influence of biofilms on porous media hydrodynamics. In: Vafai, K. (Ed.), Porous Media: Application in Biological Systems and Biotechnology. CRC press Taylor Francis Group, New York, USA, pp. 173–230.

550 551 552 553

558 Q8

C

548 549

E

546 547

R

544 545

R

542 543

N C O

540 541

U

534 535

F

556

532 533

O

554 555

We have developed methods to nondestructively evaluate the impact of microbial biofilm formation on fluid flow through a porous medium in real-time by using bioluminescence. Using this new experimental approach, it was possible to develop a relative hydraulic conductivity curve as a function of biofilm saturation, which is a new and significant advancement. The experimentally derived power law relationship between these two parameters was found to have an exponent that was significantly greater than that predicted by theoretical models, indicating a higher than expected degree of non-linearity between biofilm saturation and hydraulic conductivity. Access to such information can help researchers to better understand, and possibly manipulate, interactions between biofilm evolution, fluid flow, and chemical fate and transport in porous structures. The tools provided by this study, will, therefore, enable improved assessment of how ongoing biofilm formation affects fluid flow in porous media, which in turn, will facilitate a higher degree of control over industrially relevant biological processes such as biofiltration and bioremediation.

530 531

R O

539

529

P

4. Conclusions

527 528

Heersink, J., 2003. Basic biofilm analytical methods. In: Hamilton, M., Heersink, J., Buckingham-Meyer, K., Goeres, D. (Eds.), The Biofilm Laboratory Step-by-Step Protocols for Experimental Design, Analysis, and Data Interpretation. Cytergy Publishing Bozeman, Montana, USA, pp. 16–23. Ives, K.J., Pienvichitr, V., 1965. Kinetics of the filtration of dilute suspensions. Chem. Eng. Sci. 20, 965–973. Kelly, C.J., Hsiung, C.J., Lajoie, C.A., 2003. Kinetic analysis of bacterial bioluminescence. Biotechnol. Bioeng. 81, 370–378. Kim, Y.S., Whittle, A.J., 2006. Filtration in a porous granular medium: 1. Simulation of pore-scale particle deposition and clogging. Transp. Porous Media 65, 53–87. Kim, G., Lee, S., Kim, Y., 2006. Subsurface biobarrier formation by microorganism injection for contaminant plume control. J. Biosci. Bioeng. 101, 142–148. King, J.M.H., DiGrazia, P.M., Applegate, B., Burlage, R., Sanseverino, J., Dunbar, P., Larimer, F., Sayler, G.S., 1990. Bioluminescent reporter plasmid for naphthalene exposure and biodegradation. Science 249, 778–781. Knapp, R.M., Civan, F., McInerney, M.J., 1988. Modeling growth and transport of microorganisms in porous formations. In: Vichnevetsky, R., Borne, P., Vignes, J. (Eds.), 12th IMACS World Congress on Scientific Computation. IMACS Paris, France, pp. 676–679. Lappin-Scott, H.M., Cusack, F., MacLeod, F.A., Costerton, J.W., 1988. Nutrient resuscitation and growth of starved cells in sandstone cores: a novel approach to enhanced oil recovery. Appl. Environ. Microbiol. 54, 1373–1382. Madsen, E.L., 1991. Determining in situ biodegradation. Environ. Sci. Technol. 25, 1662–1673. Meighen, E.A., Dunlap, P.V., 1993. Physiological, biochemical and genetic control of bacterial bioluminescence. Adv. Microb. Physiol. 34, 1–67. Mitchell, A.C., Phillips, A.J., Hiebert, R., Gerlach, R., Spangler, L.H., Cunningham, A.B., 2009. Biofilm enhanced geologic sequestration of supercritical CO2. Int. J. Greenh. Gas Control 3, 90–99. Nicolella, C., van Loosdrecht, M., Heijnen, J., 2000. Wastewater treatment with particulate biofilm reactors. J. Biotechnol. 80, 1–33. Niemet, M.R., Selker, J.S., 2001. A new method for quantification of liquid saturation in 2D translucent porous media systems using light transmission. Adv. Water Resour. 24, 651–666. Okubo, T., Matsumoto, J., 1979. Effect of infiltration rate in biological clogging and water quality changes during artificial recharge. Water Resour. Res. 15, 1536–1542. Oostrom, M., Hofstee, C., Dane, J.H., Lenhard, R.J., 1998. Singlesource gamma radiation procedures for improved calibration and measurements in porous medium systems. Soil Sci. 163, 646–656. Otsu, N., 1979. A threshold selection method from gray-level histograms. IEEE Trans. Syst. Man Cybern. 9, 62–66. Ripp, S., Nivens, D.E., Ahn, Y., Werner, C., Jarrell, J., Easter, J.P., Cox, C.D., Burlage, R.S., Sayler, G.S., 2000. Controlled field release of a bioluminescent genetically engineered microorganism for bioremediation process monitoring and control. Environ. Sci. Technol. 34, 846–853. Seki, K., Thullner, M., Miyazaki, T., 2006. Moderate bioclogging leading to preferential flow paths in biobarriers. Ground Water Monit. Remediat. 26, 68–76. Seymour, J.D., Codd, S.L., Gjersing, E.L., Stewart, P.S., 2004. Magnetic resonance microscopy of biofilm structure and impact on transport in a capillary bioreactor. J Magn. Reson. 167, 322–327. Sharp, R.R., Stoodley, P., Adgie, M., Gerlach, R., Cunningham, A.B., 2005. Visualization and characterization of dynamic patterns of flow, growth and activity of biofilms growing in porous media. Water Sci. Technol. 52, 85–90. Shaw, J.J., Dane, F., Geiger, D., Kloepper, J.W., 1992. Use of bioluminescence for detection of genetically engineered microorganisms released into the environment. Appl. Environ. Microbiol. 58, 267–273. Singh, R., Paul, D., Jain, R.K., 2006. Biofilms: implications in bioremediation. Trends Microbiol. 14, 389–397. Stoodley, P., Boyle, J.D., Dodds, I., Lappin-Scott, H.M., 1998. Influence of hydrodynamics and nutrients on biofilm structure. J. Appl. Microbiol. Symp. Ser. 85, 19S–28S. Taylor, S.W., Jaffé, P.R., 1990. Biofilm growth and the related changes in the physical properties of a porous medium 1. Experimental investigations. Water Resour. Res. 26, 2153–2159. Thomas, J.M., Ward, C.H., 1989. In situ biorestoration of organic contaminants in the subsurface. Environ. Sci. Technol. 23, 760–766. Thullner, M., Zeyer, J., Kinzelbach, W., 2002. Influence of microbial growth on hydraulic properties of pore networks. Transp. Porous Media 49, 99–122. Thullner, M., Schroth, M.H., Zeyer, J., Kinzelbach, W., 2004. Modeling of a microbial growth experiment with bioclogging in a two-dimensional saturated porous media flow field. J. Contam. Hydrol. 70, 37–62. Uesugi, S.L., Yarwood, R.R., Selker, J.S., Bottomley, P.J., 2001. A model that uses the induction phase of lux-gene dependent bioluminescence in Pseudomonas fluorescens HK44 to quantify cell density in translucent porous media. J. Microbiol. Meth. 47, 317–324. van Loosdrecht, M.C., Lyklema, J., Norde, W., Zehnder, A.J.B., 1990. Influence of interfaces on microbial activity. Microbiol. Rev. 54, 75–87. Vandevivere, P., Baveye, P., Sanchez de Lozada, D., DeLeo, P., 1995. Microbial clogging of saturated soils and aquifer materials: evaluation of mathematical models. Water Resour. Res. 31, 2173–2180. Webb, O.F., Bienkowski, P.R., Matrubutham, U., Evans, F.A., Heitzer, A., Sayler, G.S., 1997. Kinetics and response of a Psuedomonas fluorescence HK44 biosensor. Biotechnol. Bioeng. 54, 491–502. West, G.B., Brown, J.H., 2005. The origin of allometric scaling laws in biology from genomes to ecosystems: towards a quantitative unifying theory of biological structure and organization. J. Exp. Biol. 208, 1575–1592.

D

538

525 526

T

536 537

conductivity follow theoretical predictions. As shown in Fig. 7b, the Clement model overestimates hydraulic conductivities at similar biofilm saturations. This can be related in part to the ignored pore throat (i.e. pore connections) clogging in the Clement model which has been shown to have a significant impact on effective hydraulic conductivity (Thullner et al., 2002). Although the experimental data were not consistent with the Clement model, it was found that an expression equivalent to Eq. (4), with n = 6.33, could be used to predict interactions between hydraulic conductivity and biofilm saturation. Higher exponent values in the bioclogging model (Eq. (4)) indicate that even small changes in porosity (i.e. low biofilm saturation) can result in severe hydraulic conductivity reductions. Such reductions in porosity could occur due to clogging of tiny pore throats, which are susceptible to plugging, and can be effectively occluded by biofilm in a manner that is not readily reversible.

E

523 524

9

Please cite this article as: Bozorg, A., et al., Using bacterial bioluminescence to evaluate the impact of biofilm on porous media hydraulic properties, J. Microbiol. Methods (2014), http://dx.doi.org/10.1016/j.mimet.2014.11.015

590 591 592 593 594 595 596 597 598 599 600 601 602 603 604 605 606 607 608 609 610 611 612 613 614 615 616 617 618 619 620 621 622 623 624 625 626 627 628 629 630 631 632 633 634 635 636 637 638 639 640 641 642 643 644 645 646 647 648 649 650 651 652 653 654 655 656 657 658 659 660 661 662 663 664 665 666 667 668 669 670 671