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\begin{document}

\title{Gas Phase Degradation of VOCs Using Supported Bacteria Biofilms}



\author[1]{Chao Xu}%
\author[1]{Ben Frigo vaz}%
\author[1]{Joshua Goering}%
\author[1]{Ping Wang}%
\affil[1]{University of Minnesota Department of Bioproducts and Biosystems Engineering}%


\vspace{-1em}



  
  \date{May 18, 2024}


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\selectlanguage{english}
\begin{abstract}
Herein we report the use of \emph{Pseudomonas putida} F1 biofilms grown
on carbonized cellulosic fibers to achieve biodegradation of airborne
VOCs in the absence of any bulk aqueous phase media. It is believed that
direct exposure of gaseous VOC substrates to biomass may eliminate
aqueous phase mass transfer resistance and facilitate VOC capture and
degradation. When tested with toluene vapor as a model VOC, the
supported biofilm could grow optimally at 300 ppm toluene and 80\%
relative humidity, with a specific growth rate of 0.425 day
\textsuperscript{-1}. During long-term VOC biodegradation tests in a
tubular packed bed reactor, biofilms achieved a toluene degradation rate
of 2.5 mg g \textsubscript{DCW} \textsuperscript{-1} h
\textsuperscript{-1} during the initial exponential growth phase.
Interestingly, the \emph{P. putida} F1 film kept biodegrading activity
even at the subsequent stationary non-growth phase. The supported
biofilms with a biomass loading of 20\% (wt) could degrade toluene at a
rate of 1.9 mg g \textsubscript{DCW} \textsuperscript{-1} h
\textsuperscript{-1} during the stationary phase, releasing CO
\textsubscript{2} at a rate of 6.4 mg g \textsubscript{DCW}
\textsuperscript{-1} h \textsuperscript{-1} at the same time (indicating
100\% conversion of substrate carbon to CO \textsubscript{2}). All the
specific degradation rates are much higher than what can be gleaned from
previously reported work. It also demonstrates the feasibility of
biofilm growth and direct gas phase degradation of VOCs without
requiring any bulk aqueous phase.%
\end{abstract}%



\sloppy


\textbf{1. Introduction}

Volatile organic compounds (VOCs) constitute a group of organic
chemicals having low T\textsubscript{b} (generally less than 250) and
have been concerned widely for adverse impacts on environment and human
health (McDonald et al., 2018; Sheu et al., 2020). Due to rapid
urbanization and industrialization, there has been a rapid increase of
VOC emission within the last century (He et al., 2019). Globally, VOC
emissions contribute significantly to environmental pollutions and
climate changes, generating photochemical smog and destroying
stratospheric ozone. Exposure to VOCs can also cause severe health
consequences (Almomani et al., 2021; Bravo et al., 2017; Hein et al.,
2018; Mat\selectlanguage{polish}ě\selectlanguage{english}jov\selectlanguage{ngerman}á et al., 2013). Short-term exposure to VOCs can cause
headaches, dizziness, fatigue, nausea, and respiratory irritation, while
long-term exposure can lead to damages to the kidney, liver, and central
nervous system (Rumchev, 2004; Wu et al., 2007; Yang et al., 2020).
VOCs, especially aromatic compounds, are often strongly recalcitrant to
biodegradation in environment. A series of restrict legislations and
standards have been formulated for VOC emission control worldwide;
however, VOCs generated in manufacturing, agriculture and transportation
industries still account for a large portion of gaseous pollutants
(Schiavon et al., 2017). To date, a variety of technologies have been
proposed for VOC treatment which can be categorized into two groups: VOC
recovery (sorption, condensation, and membrane separation) and VOC
degradation (incineration, catalytic oxidation, biodegradation,
photooxidation, and non-thermal plasma oxidation) (He et al., 2019).
Among these technologies, biodegradation is particularly appealing due
to its environmentally-benign nature and high energy efficiency over
other physical or chemical technologies (Estrada et al., 2015).

Previous research had demonstrated the feasibility of using conventional
bioreactors that typically involved microorganisms hosted in liquid
media for VOC degradation, in form of biofilters, biotrickling reactors,
and bioscrubbers (Mudliar et al., 2010; Delhoménie \& Heitz, 2005;
Detchanamurthy \& Gostomski, 2012). However, most VOCs are hydrophobic
with very limited solubility and suffer significant mass transfer
resistance in aqueous media (Cheng et al., 2016; Khan et al., 2018).
Accordingly, various bioreactor designs have been suggested and examined
to intensify VOC biodegradation by promoting substrate and biomass
interactions. That included the use of organic solvents to form
two-phase partitioning bioreactors. To our opinion, these organic phases
(including silicone oil, hexadecane, and polymeric compounds) can help
with the removal of VOC from gas phase, but do not necessary improve VOC
solubility in aqueous media hosting the biomass, as they usually offer
only limited partitioning coefficients between the two phases (Hernández
et al., 2012; Muñoz et al., 2012; Muñoz et al., 2007). Another common
approach is the use of membrane bioreactors in which porous membranes
serve as contacting surfaces interfacing gas, liquid media and biomass
for improved VOC solvation and degradation kinetics. However, the
clogging and the high cost of the membrane supports generally limited
large-scale applications of membrane bioreactors (Lebrero et al., 2013;
Reij et al., 1998). An alternative strategy is to pretreat the VOCs and
break them down to more soluble chemicals that are more vulnerable to
biodegradation. Pretreatment technologies such as UV photooxidation and
non-thermal plasma treatment have successfully been coupled with
conventional bioreactors (Almomani et al., 2021; Schiavon et al., 2015).
Such pretreatment processes may suffer from formation of toxic
byproducts, high cost and complications in scaleup operations.

As far as mass transfer resistance concerned, VOC biodegradation may
implicate gas phase diffusion, bulk liquid phase transfer, and cell
membrane adsorption and diffusion. The liquid phase mass transfer
resistance is often regarded the primary limiting factor (Mudliar et
al., 2010; Delhoménie \& Heitz, 2005; Detchanamurthy \& Gostomski, 2012;
Cheng et al., 2016). Therefore, we assume here that elimination of the
bulk liquid phase would substantially improve biodegradation efficiency.
The aim of this work is therefore to examine the feasibility of growing
microbial biofilms on solid supports without any bulk liquid phase for
VOC biodegradation. \emph{P. putida} F1 and toluene are selected as the
model bacterium and VOC, respectively. Factors regulating the growth of
biofilm on the solid support and VOC degradation efficiency are
examined. The long-term operational stability of the supported biofilm
is also demonstrated in a tubular packed bed reactor.

\textbf{2. Materials and Methods}

2.1 Media and Chemicals

Minimalist salt medium (MSM) used for producing \emph{P. putida} F1 was
prepared in our lab. MSM solution lacking carbon source substrate was
made by dissolving chemicals below to a solution of final concentrations
as: NaNO\textsubscript{3} (1 g L\textsuperscript{-1}),
KH\textsubscript{2}PO\textsubscript{4} (1 g L\textsuperscript{-1}),
K\textsubscript{2}HPO\textsubscript{4} (1 g L\textsuperscript{-1}),
MgSO\textsubscript{4} (0.2 g L\textsuperscript{-1}), KCl (0.7 g
L\textsuperscript{-1}), NaCl (0.3 g L\textsuperscript{-1}),
CaCl\textsubscript{2} (0.02 g L\textsuperscript{-1}), EDTA (2 mg
L\textsuperscript{-1}), FeCl\textsubscript{2}·4H\textsubscript{2}O (1.5
mg L\textsuperscript{-1}), H\textsubscript{3}BO\textsubscript{3} (0.06
mg L\textsuperscript{-1}), MnCl\textsubscript{2}·4H\textsubscript{2}O
(0.1 mg L\textsuperscript{-1}),
CoCl\textsubscript{2}·2H\textsubscript{2}O (0.12 mg
L\textsuperscript{-1}), ZnCl\textsubscript{2} (0.07 mg
L\textsuperscript{-1}), NiCl\textsubscript{2}·6H\textsubscript{2}O
(0.025 mg L\textsuperscript{-1}),
CuSO\textsubscript{4}·5H\textsubscript{2}O (0.02 mg
L\textsuperscript{-1}) and NaMoO\textsubscript{4}·2H\textsubscript{2}O
(0.025 mg L\textsuperscript{-1}). For MSM solid plate, extra agar (15 g
L\textsuperscript{-1}) was added along with the above chemicals. For
concentrated MSM, the concentration of the chemicals dissolved was
doubled as MSM solution. Luria-Bertani (LB) media for producing \emph{P.
putida} F1 culture to anchor biomass onto cellulosic carbon fibers
consisted of NaCl (10 g L\textsuperscript{-1}), peptone (10 g
L\textsuperscript{-1}), and yeast extract (5 g L\textsuperscript{-1}).
Toluene was purchased from Fisher Scientific (Waltham, MA). All the
other chemicals were purchased either from Fisher Scientific or
Sigma-Aldrich (St. Louis, MO).

2.2 Strain and Cultivation

\emph{Pseudomonas putida} (Trevisan) Migula (ATCC\selectlanguage{english} 700007TM) was
selected as a model bacterium in this study due to its ability of
consuming toluene (Zylstra et al., 1988). The strain was preserved in a
-70 degC refrigerator. To reactivate \emph{P. putida} F1, a 2 L glass
bottle containing 200 mL of MSM was inoculated with 1 mL of preserved
strain and capped with a Teflon-lined silicon septum having a glass bulb
with a small hole penetrating through. Toluene vapor was supplied
continuously through the small hole on the glass bulb as the sole energy
and carbon source (Claus \& Walker, 1964). The bottle was then incubated
in a shaker having 150 rpm and 30 \selectlanguage{english} under aseptic conditions for 24
hours. For further use, the activated \emph{P. putida} F1 was inoculated
onto an MSM solid plate and stored in a 4 degC refrigerator.

2.3 Preparation of Carbonized Cellulosic Fiber (CCF)

Woven gauze sponges (10 x 10 cm, 100 \% cotton) were selected as the
cellulosic material for carbonization as it has uniform structures and
high purity and requires no pretreatments. It was later carbonized in a
high-temperature tubular furnace (Sentro Tech., Model STT-1200-3.5-12,
Cleveland, OH). A typical cellulosic carbon fiber (CCF) production
process was described in our previous work (Wang et al., 2021). In
summary, gauze sponges were placed in the high-temperature tubular
furnace and then heated up to 250 \selectlanguage{english} for 3 hours with argon fed to the
furnace at a flow rate of 500 mL/min. The temperature of the furnace was
then increased to 650 \selectlanguage{english} for 4 hours and the feed was changed to a
hydrogen and argon gas mixture using flow rates of 150 mL/min and 450
mL/min, respectively. Subsequently, the furnace was further heated up to
850 \selectlanguage{english} and was held for 30 min under argon only. The chamber was finally
cooled down to room temperature at the end of the process. The strips of
CCF were then removed from the furnace and weighted. The weights were
adjusted to 0.17 g by removing excess CCF if needed.

2.4 Analytical Methods

For GC analysis of toluene and CO\textsubscript{2}, the concentrations
of toluene and CO\textsubscript{2} were analyzed in an Agilent 6890N Gas
Chromatograph equipped with FID and TCD detectors (Agilant.inc.; Santa
Clara, CA). A capillary HP-PLOT/Q column (Agilent, Santa Clara, CA) with
a length of 30 m and a diameter of 0.530 mm was used for toluene and
CO\textsubscript{2} determination. For toluene analysis, the
temperatures of the oven, injector, and FID detector were maintained at
60 degC, 220 degC, and 260 degC, respectively. For
CO\textsubscript{2}analysis, the temperatures of the oven, injector, and
TCD detector were maintained at 60 degC, 220 degC, and 250 degC, respectively.

Scanning electron microscopy (SEM) images of the CCF-supported biofilms
were taken with a HITACHI S3500N microscope (Hitachi High Technologies
America, Inc.; Schaumberg, Illinois), using 5-10 kV accelerating
voltage. Sub-milligram selections of CCF were taken from samples. The
sub-samples were then fixed, dried, and dehydrated according to
reference (Hazrin-Chong \& Manefield, 2012), after which they were
coated with 2.5 nm of gold-platinum by the mean of a Cressington 108
Auto sputter coater (Watford, England).

2.5 Preloading \emph{P. putida} F1 on CCF

MSM media was inoculated with \emph{P. putida} F1 and incubated
overnight. The OD of the culture was then adjusted to 0.5 for later use.
For preloading \emph{P. putida} F1 onto the CCF, five 50 mL centrifuge
tubes containing 0.17 g of the CCF were autoclaved and then inoculated
with 40 mL of the culture. Then four of the tubes were supplemented with
either 2 mM sucrose, 2 mM cystine, 0.4 \% (wt/v) citric acid, and 0.5 \%
(wt/v) NaCl (Chang et al., 2007; Dahlstrom et al., 2018). Finally, the
tubes were statically grown (without shaking) for 48 hours at 30 \selectlanguage{english} for
biofilm formation.

LB media was inoculated with \emph{P. putida} F1 and incubated
overnight. The OD of the culture was then adjusted to 0.5, 1.0, and 5.0,
respectively, for later use. For anchoring \emph{P. putida} F1 onto the
CCF, four 50 mL centrifuge tubes containing 0.17 g of the CCF were
autoclaved. Then three of the tubes were inoculated with 40 mL of one of
the three LB based cultures. Finally, the tubes were statically grown
(without shaking) for 48 hours at 30 \selectlanguage{english} for biofilm formation.

For \emph{P. putida} F1 biomass density analysis, biomass concentrations
in liquid phase were determined by measuring the turbidity of 1 ml of
samples of media at 600 nm with a Varian Cary(r) 50 UV-Vis
Spectrophotometer (Varian, Inc.; Lake Forest, CA). The two values were
correlated using a regression line that was made with standards
of\emph{P. putida} F1 during log-phase growth which were diluted to
specific turbidity readings and then dried at 90 \selectlanguage{english} in glass tubes
overnight and weighed. A linear fit with an R\textsuperscript{2} of 99.9
\% was achieved with triple batches and found a 0.435 optical density
correlated to 1 mg ml\textsuperscript{-1} of dry cell weight between an
optical density of 0.2 to 1.

Biomass loading of supported \emph{P. putida} F1 biofilms was obtained
by monitoring dry weight of the samples. After being treated with either
the MCM culture or the LB culture, the samples were removed from the
centrifuge tubes and washed twice using deionized water. The samples
were then placed under 90 \selectlanguage{english} in an oven for overnight drying before being
further conditioned at room temperature for 30 min. The dry weight
of\emph{P. putida} F1 on CCF was calculated based on mass balance using
dry weight data of samples before and after preloading.

2.6 Gas-phase \emph{P. putida} F1 growth in Stationary State

The testing chamber was built using an incubator (Barnstead Lab-Line,
Inc.; Melrose Park, Illinois) which allows tubing connections. Toluene
vapor was supplied into the testing chamber through the tubing and the
waste gas was removed on another side. The testing chamber was also
equipped with a humidifier (Great Innovations, LLC.; Miramar, Florida)
that was used for adjusting the humidity. To verify the reliability of
the testing chamber, two rounds of toluene concentration tests were
conducted, and each round lasted for at least five consecutive days. For
monitoring the toluene concentration, air samples were collected from
the testing chamber and then analyzed via GC-FID/TCD.

For testing of effect of humidity, five different tests were conducted
with relative humidifies (RH) of 40, 50, 60, 70, and 80 \%. Each test
lasted for five consecutive days, and the toluene concentration was set
as 60 ppm. Before putting \emph{P. putida} F1 loaded CCFs the into the
testing chamber, each sample was washed with MSM media twice and then
left for a short duration until no droplets came from the surface. For
each test, six sets of \emph{P. putida} F1 loaded CCFs were prepared,
containing a set of control samples. Each set had three pieces of the
well-prepared samples. A set of samples were removed from the testing
chamber every day for monitoring the biomass growth.

For evaluation of toxicity of toluene substrate, two different tests
were conducted with toluene concentrations of 300 and 1200 ppm. Each
test lasted for five consecutive days, and the RH was set at 80 \%.
Before putting \emph{P. putida} F1 loaded CCFs into the testing chamber,
each sample was washed with MSM media twice and then drained briefly
until no droplets came from the surface. For each test, six sets
of\emph{P. putida} F1 loaded CCFs were prepared, containing a set of
control samples. And each set have three pieces of the well-prepared
samples. A set of samples was removed from the testing chamber every day
for monitoring the biomass growth.

For biofilm stability test, RH and toluene concentration were set as
80\% and 60 ppm, respectively. Sample preparation and cultivation
protocols were the same as described above unless specified otherwise.
For toluene toxicity test, the toluene was only supplied for 6 hours
every single day to avoid excess toluene accumulation. For nutrition
limitation test, samples were washed by the concentrated MSM media
solution for providing the biofilms with more nutrition.

2.7 Long-Term Biodegradation Using Tubular Packed Bed Reactor

For tests with tubular packed bed reactor (TPBR), three pieces \emph{P.
putida} F1 loaded CCFs were produced and then placed into a tubular
packed bed reactor. The CCF-biofilm was washed with MSM media twice
before being loaded. The TPBR was constructed using 100 ml glass column
connected with air flow of 45 +- 5 ml/minute containing toluene
contaminated air of 80 +- 5 ppm and RH of 80 \% at 30 \selectlanguage{english}. Before putting
in samples, the TPBR was set for pre-running for 72 hours until the
toluene concentration and RH became stable. During 240-hour operation of
TPBR, influent and effluent toluene and
CO\textsubscript{2}concentrations were monitored with GC-FID/TCD as
described above.

\textbf{3. Results and Discussion}

3.1 Factors Affecting \emph{P. putida} F1 Biomass Preloading on CCF

Carbonized carbon fiber was chosen in this study as biofilm support
considering it has large specific surface area and hydrophobic surface
for hydrophobic VOC adsorption (Wang et al., 2021). In addition,
hydrophobic surfaces are also considered favorable for biofilm formation
as well (Farber et al., 2019). However, preloading \emph{P. putida} F1
onto CCF have been proven challenging. It has been reported that
different carbon sources might affect the initial biofilm formation on
material surface (Dahlstrom et al., 2018). Accordingly, three types of
carbon sources were tested. As can be seen from Table 1 that the biomass
of \emph{P. putida} F1 preloaded on CCF showed large differences when
treated in MCM-based culture containing different carbon sources.
Without any carbon source (thus no cell growth), the preloaded biomass
(via adsorption) is 18 +- 5 mg\textsubscript{DCW}
g\textsuperscript{-1}(Dry cell weight per gram of CCF). However, in the
presence of three different carbon sources into the MSM-based culture,
the preloading of\emph{P. putida} F1 reached 35 +- 3 (Citric acid), 73 +-
6 (Sucrose), and 29 +- 4 mg\textsubscript{DCW} g\textsuperscript{-1}
(Cystine), respectively. Previous research has also shown that alginate
produced by\emph{P. putida} F1 could promote biofilm formation (Chang et
al., 2007). NaCl was selected as alginate production stimulant. However,
that affected adversely the preloading of biomass, indicating the
alginate might not necessarily promote affinity interactions between the
CCF and cells.

Overall, sucrose promoted the highest preloading of \emph{P. putida} F1
on CCF. We also observed that \emph{P. putida} F1 could grow faster with
sucrose than other carbon sources. So the higher preloading with sucrose
could be a result of partitioning effect between the liquid media and
CCF support. Apparently, higher solution phase cellular concentration
will drive more cell assembling on CCF. We further tested this
consideration with LB culture. As a full nutrient medium, LB media can
provide \emph{P. putida} F1 with sufficient nutrition and energy
sources. To test the effect of cell concentration, LB-based culturing
was controlled with a broader cell concentration range (with OD varied
from 0.5 to 5.0). As sown in Table 2, cell concentration is positively
correlated with the preloading of biomass. With OD 5.0, the preload was
as high as 121 +- 8 mg\textsubscript{DCW} g\textsuperscript{-1}. Biomass
preloading was then conducted with OD 5.0 culturing in subsequent sample
preparations in this work.

3.2 Effect of humidity on the growth of CCF-supported biofilm

Water is essential to cell growth. In the absence of a bulk liquid phase
media, cell growth has to rely on moisture absorption from the gas
phase. Tests were conducted to examine the growth pattern of supported
biofilm under different relative humidity (RH). Figure 1A illustrates
the growth curve over a five-day period with 80 \% RH and 60 ppm
toluene. After a one-day lag phase, the sample entered an exponential
phase with a specific growth rate of 0.149 day\textsuperscript{-1} and
finally had a total biomass gain of 73
mg\textsubscript{DCW}g\textsuperscript{-1}. Decreased RH resulted in
reduced growth and less biomass accumulation. Figure
1B\textasciitilde{}E illustrate the growth with 70, 60, 50 and 40\% RH,
respectively. At 70\% RH, the samples experienced an extended lag period
before the growth phase was reached. Under this condition, the biomass
has a specific growth rate of 0.100 day\textsuperscript{-1} during the
exponential phase and finally final biomass gain was 33
mg\textsubscript{DCW} g\textsuperscript{-1}, which is substantially less
compared to 80\% RH. Decreasing the humidity further resulted in smaller
specific growth rates during the exponential growth phase, which are
0.103 day\textsuperscript{-1} at 60\% RH, 0.054 day\textsuperscript{-1}
at 50\% RH, and 0.062 day\textsuperscript{-1} at 40\% RH. Besides, the
corresponding total biomass gains were 33, 28, and 28
mg\textsubscript{DCW} g\textsuperscript{-1}, respectively. These results
indicate that water supply regulates the growth rate of \emph{P. putida}
F1. Similar RH dependency was also reported by Jin et al in a study of
supported fungal film prepared for \selectlanguage{greek}α-\selectlanguage{english}pinene treatment, which showed an
optimal RH of 85 \% (Jin et al., 2007).

SEM images also verified the growth of \emph{P. putida} F1 on CCF.
Figure 2 shows the SEM images taken for different biofilm preparations.
As shown in Figure 2B, the sporadically anchored \emph{P. putida} F1
cells can be observed on the surface of the CCF, while the surface of
untreated samples remained smooth and clean (Fig. 2A). Biofilm growth
under 80 \% RH and 60 ppm toluene showed more mature biofilm development
which formed a good coverage of the CCF support (Fig 2 C and D).
Especially, Fig. 2D shows the texture of extracellular polymeric
substances (EPS) (Nguyen et al., 2014) grown under the gaseous phase
growth mode.

3.3 Effect of toluene concentration on the growth of CCF-supported
biofilm

Theoretically, higher substrate concentration can promote faster
biodegradation kinetics before substrate saturation is reached. However,
biodegrading microbes were believed suitable for culturing at low VOC
concentrations (\textless{} 1000 ppm) due to VOC toxicity
(Gonz\selectlanguage{ngerman}ález-Martín et al., 2021). We examined biofilm growth with gaseous
VOC concentrations ranged from 60 to 1200 ppm. Compared to growth with
60 ppm toluene (Fig. 1A), when the toluene concentration was increased
to 300 and to 1200 ppm showed faster growth rate (Fig. 3). As shown in
Figure 3, no apparent lag phase could be observed, and the specific
growth rate was calculated as 0.425 day\textsuperscript{-1} at 300 ppm
toluene (Fig. 3A). The growth rate at 1200 ppm decreased from that to
0.348 day\textsuperscript{-1}. The final biomass gains for the two tests
were 72 and 65 mg\textsubscript{DCW} g\textsuperscript{-1} for 300 and
1200 ppm toluene concentration, respectively. These results showed that
increasing toluene concentration above 1000 ppm might inhibited the
biomass growth, agreeing with what reported previously with aqueous
culturing studies (Alagappan \& Cowan, 2003; Choi et al., 2008; Reardon
et al., 2000).

A comparison of growth kinetics of were \emph{P. putida} F1 observed for
different cultivation modes is presented in Table 3. Compared to our
optimal specific growth rate of 0.425 day\textsuperscript{-1}, the
specific growth rate of \emph{P. putida} F1 with same toluene
concentration (300 ppm in liquid phase) were estimated as 1.52 and 2.12
day\textsuperscript{-1}, when analyzed using Monod model and Andrews
model, respectively (Abuhamed et al., 2004; Reardon et al., 2000). It is
noteworthy that it is difficult to compare directly between gaseous and
liquid phase growth modes though, considering differences in substrate
concentration (300ppm gas phase toluene corresponds to
\textasciitilde{}80 ppm in water at sorption equilibrium). Nevertheless,
we may conclude that the 0.425 day\textsuperscript{-1} specific growth
rate is comparable to liquid phase growth kinetics (if estimated for 80
ppm using Monod model and data summarized in Table 3).

3.4 Long-term Toluene Biodegradation Potentials

To exam long-term biodegradation potentials, a tubular packed bed
reactor (TPBR) with a continuous toluene feeding stream was built.
Before packing supported biofilms in the TPBR, the reactor was tested
for 24 hours to achieve a stable operation condition. The operation
condition set for TPBR was 80 \% RH, with a gas feeding rate fixed at 45
ml/min. Toluene biodegradation rate and CO\textsubscript{2} production
rate were monitored during the tests (Figure 4). The biomass loadings
were also measured before and after the long-term operation. Compared
with the initial biomass loading of 163
mg\textsubscript{DCW}g\textsuperscript{-1}, the final biomass was
measured as 203 mg\textsubscript{DCW} g\textsuperscript{-1} on CCF, and
the final biomass gain was 40 mg (about 25\% mass gain) over 10 days. A
total toluene consumed of 47 mg. Therefore, the yield coefficient of
biomass was calculated as 0.85 g/g, which agrees well with what have
been previously reported (ranged 0.58 \textasciitilde{} 1.28 g/g)
(Abuhamed et al., 2004; Reardon et al., 2000).

At the beginning of the operation (0 \textasciitilde{} 12 h), the
biofilm experienced a lag phase. After that, apparent degradation of
toluene took place (as reflected from the difference between the inlet
and outlet toluene concentration, 12\textasciitilde{}36 h, Fig. 4). At
the same time, the outlet CO\textsubscript{2} increased from 202 ppm to
369 ppm. Degradation proceeded after 36 h but with a reduced reaction
rate, as indicated by the narrowed gaps between feed and outlet
concentrations (both toluene and CO\textsubscript{2}). The first few
hours after the lag phase should correspond the a exponential growth
phase of the biofilm, for which toluene consumption and
CO\textsubscript{2} generation rates were calculated as 2.5 mg
g\textsubscript{DCW}\textsuperscript{-1} h\textsuperscript{-1} and 5.8
mg g\textsubscript{DCW}\textsuperscript{-1} h\textsuperscript{-1},
respectively (based on initial biomass loading of 16\%-wt). Reactions
exceeding 72 h appeared to be stable throughout the operation, and
should echo a stationary degradation phase. The inlet and outlet toluene
concentrations were around 90 ppm and 70 ppm, respectively. In the
stationary phase, supported biofilms loading increased to 20\%-wt,
achieved a degradation rate of 1.9 mg
g\textsubscript{DCW}\textsuperscript{-1} h\textsuperscript{-1} and
releasing 6.4 mg
g\textsubscript{DCW}\textsuperscript{-1}h\textsuperscript{-1}
CO\textsubscript{2} at the same time (indicating 100\% conversion of
toluene to CO\textsubscript{2}).

Overall the direct gaseous phase biodegradation showed exciting
enhancement in reaction efficiency if compared to degradation rates
reported previously by others. The growth phase degradation rate,
measured as 2.5 mg
g\textsubscript{DCW}\textsuperscript{-1}h\textsuperscript{-1} in this
work, is as high as 10-fold of what has been observed for liquid media
growth-associated biodegradation rate of toluene (ranged from 0.17 to
0.46 mg g\textsubscript{DCW}\textsuperscript{-1} h\textsuperscript{-1}
(Díaz et al., 2008; Muñoz et al., 2008). The nongrowth-associated
stationary degradation phase is a comparable reaction mode to the latex
coating film of \emph{P.putida} F1 as reported previously by Estrada et
al. (Estrada et al., 2015), which enabled degradation rates (in the rage
of 0.034 to 0.321 mg
g\textsubscript{DCW}\textsuperscript{-1}h\textsuperscript{-1}) that were
also much lower than what is observed in this work (1.9 mg
g\textsubscript{DCW}\textsuperscript{-1}h\textsuperscript{-1}).

3.4 Biofilm Growth Limiting Factors

The above observation clearly indicated that \emph{P. putida} F1 biofilm
could enable both growth-associated and non-growth associated
degradation of toluene. The higher growth-associated degradation rate
prompt us to investigate the factors that limit the duration of the
exponential growth of CCF-supported \emph{P. putida} F1 biofilm. As
mentioned earlier, toluene toxicity may inhibit biomass growth, and
there is a possibility for concentrated toluene accumulation under
continuous feed operations considering that CCF could adsorb well
toluene (Wang et al., 2021). In addition, insufficient nutrients (no
nutrition amendment during the operation) could also be limiting.

To evaluate the effect of toluene accumulation, biofilm growth was
tested in the chamber reactor under 80 \% RH, but with limited toluene
feed (supplied with 60 ppm toluene intermittently 6 h per day). In this
case, excess toluene adsorbed onto the surface of CCFs would be released
or degraded timely. As can be seen from Figure 5A, the \emph{P.
putida}F1 growth pattern did not change from growth with constant
toluene concentration (Fig. 1A). There was a lag phase followed by an
exponential phase, and finally entered a stationary phase. That process
appeared to be slower, but only slightly extended. The biomass gains
of\emph{P. putida} F1 in this test was 45
mg\textsubscript{DCW}g\textsuperscript{-1}, which was lower than 73
mg\textsubscript{DCW}g\textsuperscript{-1} obtained under optimal
condition. This can be attributed to the less amount of toluene supply
in the chamber during the growth of \emph{P. putida} F1. This result
indicates that toluene accumulation is not a factor that limits growth
period of \emph{P. putida} F1.

To test the nutrient limitation hypothesis, samples were pretreated with
concentrated MSM media. Microbial growth (Fig. 5B) experienced a longer
exponential phase and entered the stationary phase after 4 days. The
biomass gain in this test was 136
mg\textsubscript{DCW}g\textsuperscript{-1}, which was about two times
greater than that we observed under standard conditions. The much
increased biomass growth also become detectable with eyes (Fig. 6). As
can be seen in Figure 6, there were more white accumulation appearing on
doubled nutrient treated CCF than the samples normally treated. However,
since the growth duration was not apparently extended, excess nutrition
simply increased the overall growth rate.

In fact, previous research conducted by other researchers have also
indicated biofilms did not keep growing after three or four days. One
potential reason for the halt in growth could be metabolite accumulation
in biofilms, as toluene degradation with the \emph{P. putida} F1 could
accumulate a mutagenic metabolite benzyl alcohol (Bordel et al., 2007).
From our observations, this could be the primary regulating factor, but
will require more systematic investigation to confirm and eventually
extend the growth-associated biodegradation operations.

\textbf{4. Conclusion}

\emph{P. putida} F1 was successfully preloaded onto CCFs and applied for
airborne toluene degradation. Studies showed that the \emph{P. putida}F1
was able to grow on the CCF support and eventually formed mature
biofilms under feeding with gaseous toluene. The CCF-supported \emph{P.
putida} F1 biofilms can grow at the RH range from 40 to 80\% and the
biomass grow faster at higher humidity levels. The optimal specific
growth rate of \emph{P. putida} F1 in gas phase was actually comparable
to that in liquid phase. The CCF-supported \emph{P. putida} F1 enable
both growth-associated and non-growth associated biodegradation
activities. Operated without any bulk liquid medium phase, the biofilm
achieved direct degradation of gas phase VOC at rates of about one-order
of magnitude higher than what has been previously reported for liquid
culturing or immobilized cells. These results suggest that liquid phase
mass transfer is a significant limiting factor for traditional
biodegradation reactions, and biofilms operated in gas phases can
promote efficient substrate-cell interactions and thus intensifying
biotransformation reactions.

\textbf{5. Acknowledgements}

We want to thank Ellen Guo for assisting part of the experimental work.
The authors thank supports from MnDrive Environment Fund and Industrial
Partnership for Research in Interfacial and Materials Engineering
(Iprime) Fund of University of Minnesota.

\textbf{6. Reference}

Abuhamed, T., Bayraktar, E., Mehmeto\selectlanguage{polish}ğ\selectlanguage{english}lu, T., \& Mehmeto\selectlanguage{polish}ğ\selectlanguage{english}lu, \selectlanguage{ngerman}Ü. (2004).
Kinetics model for growth of Pseudomonas putida F1 during benzene,
toluene and phenol biodegradation. \emph{Process Biochemistry}
,\emph{39} (8), 983--988. https://doi.org/10.1016/S0032-9592(03)00210-3

Alagappan, G., \& Cowan, R. (2003). Substrate inhibition kinetics for
toluene and benzene degrading pure cultures and a method for collection
and analysis of respirometric data for strongly inhibited
cultures.\emph{Biotechnology and Bioengineering} , \emph{83} (7),
798--809. https://doi.org/10.1002/bit.10729

Almomani, F., Rene, E. R., Veiga, M. C., Bhosale, R. R., \& Kennes, C.
(2021). Treatment of waste gas contaminated with dichloromethane using
photocatalytic oxidation, biodegradation and their
combinations.\emph{Journal of Hazardous Materials} , \emph{405} ,
123735. https://doi.org/10.1016/j.jhazmat.2020.123735

Bordel, S., Muñoz, R., Díaz, L. F., \& Villaverde, S. (2007). New
insights on toluene biodegradation by \emph{Pseudomonas putida} F1:
Influence of pollutant concentration and excreted
metabolites.\emph{Applied Microbiology and Biotechnology} , \emph{74}
(4), 857--866. https://doi.org/10.1007/s00253-006-0724-8

Bravo, D., Ferrero, P., Penya-roja, J. M., Álvarez-Hornos, F. J., \&
Gabaldón, C. (2017). Control of VOCs from printing press air emissions
by anaerobic bioscrubber: Performance and microbial community of an
on-site pilot unit. \emph{Journal of Environmental Management}
,\emph{197} , 287--295. https://doi.org/10.1016/j.jenvman.2017.03.093

Chang, W.-S., van de Mortel, M., Nielsen, L., Nino de Guzman, G., Li,
X., \& Halverson, L. J. (2007). Alginate production by \emph{Pseudomonas
putida} creates a hydrated microenvironment and contributes to biofilm
architecture and stress tolerance under water-limiting
conditions.\emph{Journal of Bacteriology} \emph{189} (22), 8290--8299.
https://doi.org/10.1128/JB.00727-07

Cheng, Y., He, H., Yang, C., Zeng, G., Li, X., Chen, H., \& Yu, G.
(2016). Challenges and solutions for biofiltration of hydrophobic
volatile organic compounds. \emph{Biotechnology Advances} , \emph{34}
(6), 1091--1102. https://doi.org/10.1016/j.biotechadv.2016.06.007

Choi, N.-C., Choi, J.-W., Kim, S.-B., \& Kim, D.-J. (2008). Modeling of
growth kinetics for \emph{Pseudomonas putida} during toluene
degradation. \emph{Applied Microbiology and Biotechnology} ,\emph{81}
(1), 135--141. https://doi.org/10.1007/s00253-008-1650-8

Claus, D., \& Walker, N. (1964). The decomposition of toluene by soil
bacteria. \emph{Microbiology,} \emph{36} (1), 107--122.
https://doi.org/10.1099/00221287-36-1-107

Dahlstrom, K. M., Collins, A. J., Hogan, D. A., \& O'Toole, G. A.
(2018). A multimodal strategy used by a large c-di-GMP
network.\emph{Journal of Bacteriology} , \emph{200} (8), 19.
https://doi.org/10.1128/JB.00703-17

Delhoménie, M.-C., \& Heitz, M. (2005). Biofiltration of air: A
review.\emph{Critical Reviews in Biotechnology} , \emph{25} (1--2),
53--72. https://doi.org/10.1080/07388550590935814

Detchanamurthy, S., \& Gostomski, P. A. (2012). Biofiltration for
treating VOCs: An overview. \emph{Reviews in Environmental Science and
Bio/Technology} , \emph{11} (3), 231--241.
https://doi.org/10.1007/s11157-012-9288-5

Díaz, L. F., Muñoz, R., Bordel, S., \& Villaverde, S. (2008). Toluene
biodegradation by \emph{Pseudomonas putida} F1: Targeting culture
stability in long-term operation. \emph{Biodegradation} , \emph{19} (2),
197--208. https://doi.org/10.1007/s10532-007-9126-6

Estrada, J. M., Bernal, O. I., Flickinger, M. C., \& Deshusses, M. A.
(2015). Biocatalytic coatings for air pollution control: A proof of
concept study on VOC biodegradation. \emph{Biotechnology and
Bioengineering} , \emph{112} (2), 10.

Farber, R., Dabush-Busheri, I., Chaniel, G., Rozenfeld, S., Bormashenko,
E., Multanen, V., \& Cahan, R. (2019). Biofilm grown on wood waste
pretreated with cold low-pressure nitrogen plasma: Utilization for
toluene remediation. \emph{International Biodeterioration \&
Biodegradation} , \emph{139} , 62--69.
https://doi.org/10.1016/j.ibiod.2019.03.003

González-Martín, J., Kraakman, N. J. R., Pérez, C., Lebrero, R., \&
Muñoz, R. (2021). A state--of--the-art review on indoor air pollution
and strategies for indoor air pollution control. \emph{Chemosphere}
,\emph{262} , 128376. https://doi.org/10.1016/j.chemosphere.2020.128376

Hazrin-Chong, N. H., \& Manefield, M. (2012). An alternative SEM drying
method using hexamethyldisilazane (HMDS) for microbial cell attachment
studies on sub-bituminous coal. \emph{Journal of Microbiological
Methods} , \emph{90} (2), 96--99.
https://doi.org/10.1016/j.mimet.2012.04.014

He, C., Cheng, J., Zhang, X., Douthwaite, M., Pattisson, S., \& Hao, Z.
(2019). Recent advances in the catalytic oxidation of volatile organic
compounds: A review based on pollutant sorts and sources. \emph{Chemical
Reviews} , \emph{119} (7), 4471--4568.
https://doi.org/10.1021/acs.chemrev.8b00408

Hein, L., White, L., Miles, A., \& Roberts, P. (2018). Analysing the
impacts of air quality policies on ecosystem services; a case study for
Telemark, Norway. \emph{Journal of Environmental Management} ,\emph{206}
, 650--663. https://doi.org/10.1016/j.jenvman.2017.10.073

Hernández, M., Quijano, G., \& Muñoz, R. (2012). Key role of microbial
characteristics on the performance of VOC biodegradation in two-liquid
phase bioreactors. \emph{Environmental Science \& Technology} ,\emph{46}
(7), 4059--4066. https://doi.org/10.1021/es204144c

Jin, Y., Guo, L., Veiga, M. C., \& Kennes, C. (2007). Fungal
biofiltration of \selectlanguage{greek}α-\selectlanguage{english}pinene: Effects of temperature, relative humidity,
and transient loads. \emph{Biotechnology and Bioengineering} ,\emph{96}
(3), 433--443. https://doi.org/10.1002/bit.21123

Khan, A. M., Wick, L. Y., \& Thullner, M. (2018). Applying the Rayleigh
approach for stable isotope-based analysis of VOC biodegradation in
diffusion-dominated systems. \emph{Environmental Science \& Technology}
,\emph{52} (14), 7785--7795. https://doi.org/10.1021/acs.est.8b01757

Lebrero, R., Volckaert, D., P\selectlanguage{ngerman}érez, R., Muñoz, R., \& Van Langenhove, H.
(2013). A membrane bioreactor for the simultaneous treatment of acetone,
toluene, limonene and hexane at trace level concentrations. \emph{Water
Research} , \emph{47} (7), 2199--2212.
https://doi.org/10.1016/j.watres.2013.01.041

Mat\selectlanguage{polish}ě\selectlanguage{english}jov\selectlanguage{ngerman}á, L., Topka, P., Kalu\selectlanguage{polish}ž\selectlanguage{english}a, L., Pitk\selectlanguage{ngerman}äaho, S., Ojala, S., Gaálová,
J., \& Keiski, R. L. (2013). Total oxidation of dichloromethane and
ethanol over ceria--zirconia mixed oxide supported platinum and gold
catalysts. \emph{Applied Catalysis B: Environmental} , \emph{142--143} ,
54--64. https://doi.org/10.1016/j.apcatb.2013.04.069

McDonald, B. C., de Gouw, J. A., Gilman, J. B., Jathar, S. H., Akherati,
A., Cappa, C. D., Jimenez, J. L., Lee-Taylor, J., Hayes, P. L., McKeen,
S. A., Cui, Y. Y., Kim, S.-W., Gentner, D. R., Isaacman-VanWertz, G.,
Goldstein, A. H., Harley, R. A., Frost, G. J., Roberts, J. M., Ryerson,
T. B., \& Trainer, M. (2018). Volatile chemical products emerging as
largest petrochemical source of urban organic emissions. \emph{Science}
,\emph{359} (6377), 760--764. https://doi.org/10.1126/science.aaq0524

Mudliar, S., Giri, B., Padoley, K., Satpute, D., Dixit, R., Bhatt, P.,
Pandey, R., Juwarkar, A., \& Vaidya, A. (2010). Bioreactors for
treatment of VOCs and odours -- A review. \emph{Journal of Environmental
Management} , \emph{91} (5), 1039--1054.
https://doi.org/10.1016/j.jenvman.2010.01.006

Muñoz, R., Daugulis, A. J., Hernández, M., \& Quijano, G. (2012). Recent
advances in two-phase partitioning bioreactors for the treatment of
volatile organic compounds. \emph{Biotechnology Advances} , \emph{30}
(6), 1707--1720. https://doi.org/10.1016/j.biotechadv.2012.08.009

Muñoz, R., Díaz, L. F., Bordel, S., \& Villaverde, S. (2008). Response
of \emph{Pseudomonas putida} F1 cultures to fluctuating toluene loads
and operational failures in suspended growth
bioreactors.\emph{Biodegradation} , \emph{19} (6), 897--908.
https://doi.org/10.1007/s10532-008-9191-5

Muñoz, R., Villaverde, S., Guieysse, B., \& Revah, S. (2007). Two-phase
partitioning bioreactors for treatment of volatile organic
compounds.\emph{Biotechnology Advances} , \emph{25} (4), 410--422.
https://doi.org/10.1016/j.biotechadv.2007.03.005

Nguyen, P. Q., Botyanszki, Z., Tay, P. K. R., \& Joshi, N. S. (2014).
Programmable biofilm-based materials from engineered curli
nanofibres.\emph{Nature Communications} , \emph{5} (1), 4945.
https://doi.org/10.1038/ncomms5945

Reardon, K. F., Mosteller, D. C., \& Rogers, J. D. B. (2000).
Biodegradation kinetics of benzene, toluene, and phenol as single and
mixed substrates for \emph{Pseudomonas putida} F1. \emph{Biotechnology
and Bioengineering} , \emph{69} (4), 16.

Reij, M. W., Keurentjes, J. T. F., \& Hartmans, S. (1998). Membrane
bioreactors for waste gas treatment. \emph{Journal of Biotechnology}
,\emph{59} (3), 155--167. https://doi.org/10.1016/S0168-1656(97)00169-7

Rumchev, K. (2004). Association of domestic exposure to volatile organic
compounds with asthma in young children. \emph{Thorax} , \emph{59} (9),
746--751. https://doi.org/10.1136/thx.2003.013680

Schiavon, M., Scapinello, M., Tosi, P., Ragazzi, M., Torretta, V., \&
Rada, E. C. (2015). Potential of non-thermal plasmas for helping the
biodegradation of volatile organic compounds (VOCs) released by waste
management plants. \emph{Journal of Cleaner Production} , \emph{104} ,
211--219. https://doi.org/10.1016/j.jclepro.2015.05.034

Schiavon, M., Schiorlin, M., Torretta, V., Brandenburg, R., \& Ragazzi,
M. (2017). Non-thermal plasma assisting the biofiltration of volatile
organic compounds. \emph{Journal of Cleaner Production} , \emph{148} ,
498--508. https://doi.org/10.1016/j.jclepro.2017.02.008

Sheu, R., Stönner, C., Ditto, J. C., Klüpfel, T., Williams, J., \&
Gentner, D. R. (2020). Human transport of thirdhand tobacco smoke: A
prominent source of hazardous air pollutants into indoor nonsmoking
environments. \emph{Science Advances} , \emph{6} (10), eaay4109.
https://doi.org/10.1126/sciadv.aay4109

Wang, Y.-H., Bayatpour, S., Qian, X., Frigo-Vaz, B., \& Wang, P. (2021).
Activated carbon fibers via reductive carbonization of cellulosic
biomass for adsorption of nonpolar volatile organic
compounds.\emph{Colloids and Surfaces A: Physicochemical and Engineering
Aspects} ,\emph{612} , 125908.
https://doi.org/10.1016/j.colsurfa.2020.125908

Wu, F., Jacobs, D., Mitchell, C., Miller, D., \& Karol, M. H. (2007).
Improving Indoor Environmental Quality for Public Health: Impediments
and Policy Recommendations. \emph{Environmental Health Perspectives}
,\emph{115} (6), 953--957. https://doi.org/10.1289/ehp.8986

Yang, N., Wang, C., \& Han, M.-F. (2020). Gel-encapsulated
microorganisms used as a strategy to rapidly recover biofilters after
starvation interruption. \emph{Journal of Environmental Management}
,\emph{261} , 110237. https://doi.org/10.1016/j.jenvman.2020.110237

Zylstra, G. J., McCombie, W. R., Gibson, D. T., \& Finette, B. A.
(1988). Toluene degradation by \emph{Pseudomonas putida} F1: Genetic
organization of the tod operon. \emph{Applied and Environmental
Microbiology} , \emph{54} (6), 1498--1503.
https://doi.org/10.1128/aem.54.6.1498-1503.1988

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