1
Extracellular DNA promotes efficient extracellular electron transfer by pyocyanin in
Pseudomonas aeruginosa
biofilms.
Authors:
Scott H. Saunders
1
, Edmund C.M. Tse
2
, Matthew D. Yates
3
, Fernanda Jiménez
Otero
4
, Scott
A. Trammell
3
, Eric D.A. Stemp
5
, Jacqueline K. Barton
2
*, Leonard M. Tender
3
* and Dianne K.
Newman
1,6
*
Affiliations:
1
Division of Biology and Biological Engineering, Caltech, Pasadena, CA.
2
Division of Chemistry and Chemical Engineering, Caltech, Pasadena, CA.
3
Center for Bio/Molecular Science and Engineering, Naval Research Laboratory, Washington, DC.
4
George Mason University, Fairfax, VA.
5
Department of Physical Sciences, Mt. Saint Mary’s University, Los Angeles,
CA.
6
Division of Geological and Planetary Sciences, Caltech, Pasadena, CA.
*Correspondence to: jkbarton@caltech.edu,
leonard.tender@nrl.navy.mil
, dkn@caltech.edu (
DKN=
lead
contact).
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SUMMARY
:
Extracellular electron transfer (EET
), the process whereby
cells
access
electron acceptors or donors that
reside many cell lengths away,
enables metabolic activity by microorganisms, particularly under
oxidant-limited conditions that occur in multicellular bacterial biofilms. Although different mechanisms
underpin this process in select organisms, a widespread strategy involves extracellular electron shuttles,
redox-active metabolites that are secreted and recycled by diverse bacteria. How these shuttles catalyze
electron transfer within biofilms without being lost to the environment has been a long-standing
question. Here, we show that phenazine electron shuttles mediate efficient EET through interactions
with extracellular DNA (eDNA)
in
Pseudomonas aeruginosa
biofilms, which are important in nature
and disease. Retention of pyocyanin (PYO) and phenazine carboxamide in the biofilm matrix i
s
facilitated by binding to eDNA
. In vitro, different phenazines can exchange electrons in the presence or
absence of DNA and phenazines can participate directly in redox reactions through DNA; the biofilm
eDNA can also support rapid electron transfer betw
een
redox active intercalators. Electrochemical
measurements of biofilms indicate that retained PYO supports an efficient redox cycle with rapid EET
and slow loss from the biofilm. Together, these results establish that eDNA facilitat
es phenazine
metabolic processes in
P. aeruginosa
biofilms, suggesting a model for how extracellular electron
shuttles achieve retention and efficient EET in biofilms.
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INTRODUCTION
:
Microbial biofilms are ubiquitous in natural and engineered contexts, spanning plant roots to chronic
human infections to anaerobic digestors (Watnick and Kolter, 2000). As biofilms develop, metabolic
stratification occurs, driven by steep concentration gradients of substrates, such as oxygen, that are
consumed by cells at the biofilm periphery faster than the substrates can diffuse into the biofilm interior
(Stewart, 2003; Stewart and Franklin, 2008; Xu et al., 1998). Indeed, oxidant limitation is a generic
challenge for cells that inhabit biofilm microenvironments where electron donors are abundant, yet
electron acceptors are not. One widespread strategy microbes employ to overcome this challenge is to
channel electrons derived from intracellular metabolism to extracellular oxidants at a distance (Shi et al.,
2016). Known as “extracellular electron transfer” (EET), this process requires electron carriers to bridge
the gap, be they outer membrane-associated or extracellular cytochromes (Jiménez Otero et al., 2018;
Richter et al., 2009; Xu et al., 2018), cytochrome-replete “nanowires” (Subramanian et al., 2018; Wang
et al., 2019), cable bacteria conductive filaments (Cornelissen et al., 2018), or redox-active small
molecules (Glasser et al., 2017a; Hernandez and Newman, 2001). While the putative molecular
components underpinning different EET processes have been described in a variety of organisms, a
detailed understanding of how these components achieve EET remains an important research goal across
diverse systems.
In contrast to the intense study of microbial nanowires (Malvankar et al., 2011; Reguera et al., 2005;
Wang et al., 2019), less attention has been paid to how small soluble (physically diffusive) extracellular
electron shuttles facilitate EET beyond interactions at the cell surface (Light et al., 2018; Marsili et al.,
2008; Xu et al., 2016). In part, this neglect is due to the challenges involved in identifying and studying
small molecule metabolites, compared to the multiheme cytochromes observed in many genomes of
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organisms known to engage in EET. Accordingly, to study extracellular electron shuttling, we have
chosen to work with a model system that employs a relatively well studied and tractable set of shuttles
called phenazines. Phenazines are colorful redox-active molecules that are produced by numerous
microbial species including the bacterium,
Pseudomonas aeruginosa
(Turner and Messenger, 1986)
.
P.
aeruginosa
strains are ubiquitous yet perhaps most well-known for their roles in chronic infections
where their growth as biofilms renders them antibiotic tolerant and contributes to patient morbidity and
mortality (Costerton et al., 1999)
; importantly, phenazines support the development of anoxic, antibiotic
tolerant biofilm regions (Dietrich et al., 2013a; Jo et al., 2017; Schiessl et al., 2019)
. While significant
progress has been made in defining the composition of the
P. aeruginosa
biofilm matrix (Colvin et al.,
2012) and mapping the zones of phenazine production within it (Bellin et al., 2014, 2016), we still have
much to learn about how phenazines facilitate EET within the matrix.
Intriguingly, while the
P. aeruginosa
biofilm matrix comprises a heterogeneous group of polymers,
extr
acellular DNA (eDNA) from dead cells is a significant contributor
(Allesen
-Holm et al., 2006;
Whitchurch et al., 2002), accounting for the majority of the matrix polymers in some cases (Matsukawa
and Greenberg, 2004; Steinberger and Holden, 2005). Phenazines have long been known to intercalate
into double stranded DNA
in vitro
(Hollstein and Van Gemert, 1971), and more recently, it was
suggested that the phenazine pyocyanin (PYO) can participate in
DNA-mediated charge transfer (DNA
CT) chemistry
in vitro
(Das et al., 2015). Together with the observation that PYO promotes eDNA
release by stimulating cell lysis (Das and Manefield, 2012), these facts led to speculation that phenazine-
eDNA interactions might facilitate biofilm EET (Das et al., 2015). Notably, the ability of PYO to
stimulate cell lysis changes according to the environment: when cells are oxidatively stressed (
i.e.
oxidant replete but reductant limited) and ATP limited, PYO is toxic; whereas when they are reductively
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stressed (
i.e.
reductant replete but oxidant limited), PYO promotes viability and biofilm aggregate
expansion (Costa et al., 2017; Meirelles and Newman, 2018). This observation suggests the intriguing
possibility that cell lysis by a small percentage of the population early on might later promote EET once
biofilms have developed anoxic zones where extracellular electron shuttles support metabolism. Though
a variety of roles for eDNA in biofilms have been proposed, including serving as a structural support,
nutrient and/or genetic reservoir (Flemming and Wingender, 2010), to our knowledge, that eDNA may
stimulate biofilm metabolism by facilitating EET has not been tested.
The current model of the phenazine redox cycle in biofilms can be broadly defined (Fig. 1A). In anoxic
regions, oxidized phenazines are reduced intracellularly by metabolic reactions that support these cells
(Glass
er et al., 2014, 2017b; Jo et al., 2017; Wang et al., 2010)
. These reduced phenazines physically
diffuse through the extracellular matrix toward the oxic region where they react abiotically with
molecular oxygen. Upon re-oxidation, phenazines return to the anoxic region of the biofilm to complete
the redox cycle. Although studies have begun to characterize the reactions on either side of the redox
cycle, very little is known about how phenazines operate in the intervening extracellular matrix.
Theoretical studies suggest that physical diffusion of oxidized phenazine towards the biofilm interior
and reduced phenazine toward the biofilm periphery may be fast enough to support the metabolism of
the oxidant limited cells (Glasser et al., 2017a; Kempes et al., 2014). However, these studies assume a
closed system, and an unresolved paradox has been how diffusible extracellular molecules could
function in a redox cycle without being lost from the biofilm to the environment. Here we explore how
phenazine electron transfer may be reconciled with phenazine retention. Specifically, we ask: Are
phenazines retained? What mechanisms of electron transfer are compatible with phenazine retention? Is
phenazine electron transfer
in vivo
fast compared to phenazine loss? Our motivation to answer these
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questions arises not only from a desire to constrain the model of phenazine redox cycling within
P.
aeruginosa
biofilms, but more broadly, to identify a potentially generalizable strategy for how diverse
electron shuttles enable EET.
RESULTS
:
We studied three major phenazine derivatives made by
P. aeruginosa
strain UCBPP PA14 (Schroth et
al., 2018): phenazine carboxylate (PCA), phenazine carboxamide (PCN), and pyocyanin (PYO) (Fig.
1B). Beyond studying wildtype (WT) produced phenazines, we also explore the effects of individual
synthetic phenazines on a mutant that does not produce phenazines,
∆
phz
(
∆
phzA1-
G1
,
∆
phzA2-
G2
), or
on a mutant that is also incapable of modifying PCA,
∆
phz*
(
∆
phz,
∆
phzMS,
∆
phzH
). Experiments were
performed with two different types of biofilms: macroscopic colony biofilms grown on nutrient agar
surfaces (Fig. 1C, Fig. S1A) and microscopic biofilms attached to the surface of an interdigitated
microelectrode array suspended in liquid medium. Phenazine-dependent biofilm phenotypes operate
similarly at both scales (Ramos et al., 2010)
, so
we selected the biofilm cultivation method for any given
experiment based on which was best suited to answering our specific research question.
Colony biofilms retain PCN and PYO, but not PCA
First, we sought to quantify phenazine retention by colony biofilms (
Fig 1
C-D). In contrast to previous
work that used an electrode array to electrochemically measure the spatial distribution of phenazines that
physically diffuse into agar underneath colony biofilms (Bellin et al., 2014, 2016), we used liquid
chromatography-mass spectrometry (LC-MS) to quantify extracted endogenous phenazines from the
biofilms and compare their concentrations to that in the underlying agar (Fig 1D-
F). Colony biofilms
could be cleanly separated from the agar, because they were separated by a 0.2
μ
m membrane filter,
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which did not affect the results (
Fig. S1B-C). Overall, PCA, PCN, and PYO concentrations varied by
more than 10-fold in the biofilms reaching concentrations of ~15
μ
M PCA, ~ 400
μ
M PCN, and ~80
μ
M
PYO. Comparing biofilm to agar concentrations showed that PCN and PYO were enriched in the
biofilm 10-fold and 30-fold, respectively, while PCA reached similar concentrations in the biofilm and
the agar (Fig. 1E-F). This suggested that PCN and PYO were strongly retained by the biofilm and PCA
was not. Importantly, lysing resuspended biofilm cells by sonication prior to phenazine quantification
did not strongly affect the results (Fig. S1D), indicating that the measured pools of phenazines were
predominantly retained in the extracellular matrix rather than intracellularly.
To test if differential phenazine retention was caused by a spatial or temporal difference in biosynthesis,
we grew
D
phz
* colony biofilms with synthetic phenazines in the agar and quantified phenazines taken
up by the biofilm. Incubation
with
> 10
μ
M PYO resulted in >200
μ
M PYO accumulation in the biofilm
(Fig. 1G). PCN accumulated to a lesser extent, and PCA biofilm uptake was minimal (<50
μ
M) even
with 200
μ
M added to the agar (Fig. 1G).
D
phz
* colonies transferred from phenazine agar to fresh agar
after 3 days of growth retained phenazines in the same pattern as the wild type (WT) over 24 h (Fig.
1H), demonstrating that the observed phenazine retention does not depend on endogenous phenazine
production. W
ild
-type colony biofilms exhibit relatively thick and smooth morphologies that contain
deep anoxic regions that are thought to be supported by phenazines.
∆
phz*
colony biofilms exhibit
different colony morphologies that are thin and highly wrinkled, which is thought to be a physiological
adaptation to maximize surface area and oxygen penetration in the absence of phenazines as shown for
∆
phz
(Dietrich et al., 2013b). Notably, only incubation of
D
phz
* colonies with exogenous PYO
appreciably complemented the colony wrinkling phenotype (Fig. S1A).
P. aeruginosa
colony biofilms
thus appear able to take up and use significant amounts of exogenous PYO, and PCN to a lesser extent.
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These results predict that colony biofilms contain an extracellular matrix component that binds and
effectively retains PYO and PCN, but not PCA.
Phenazines differentially
bind
extracellular
DNA
The extracellular matrix in
P. aeruginosa
PA14 biofilms is known to be primarily composed of two
polymers: DNA from dead cells (eDNA) and the polysaccharide, Pel (Colvin et al., 2011; Das and
Manefield, 2012). To test the hypothesis that eDNA in the biofilm matrix was responsible for binding
phenazines, we quantified the binding affinity of oxidized PCA, PCN, and PYO for double-stranded (ds)
DNA
in vitro
using isothermal titration calorimetry (Fig. 2A). As expected, oxidized PCA showed no
detectable binding because it is negatively charged, as is the phosphate backbone of DNA at pH 7. In
contrast, oxidized PCN (K
D
= 194
μ
M; 95% C.I. 148 – 305
μ
M) and PYO (K
D
= 13
μ
M; 95% C.I. 6.5 –
49
μ
M) both bind ds DNA, and these results were consistent with ethidium bromide displacement and
microscale thermophoresis binding assays (Fig. S
2A
-B)
(Das et al., 2015). Notably, these
in vitro
phenazine-DNA binding affinities correlate with their
in vivo
retention ratio ( [biofilm] / [agar] ), where
PYO is retained in the biofilm significantly more than PCN, and PCA is not retained. Reduced
PYO
showed no change in endogenous fluorescence upon addition of calf thymus DNA (Fig. S2C)
, which is
unexpected for strong intercalative binding. Therefore, the DNA binding affinity of PYO
is likely redox
dependent.
To determine whether phenazine-eDNA binding occurs
in vivo
, we treated 3-day old WT biofilms with
DNase I for 24 hrs. These experiments were performed with DNase I spotted on tryptone agar medium
rather than its optimal buffer, as controls showed that buffer alone significantly disturbed the biofilm
(Fig. S3A-C). Despite a low activity for DNase under these conditions, DNase-treated biofilms showed
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significantly lower biofilm PCN and PYO concentrations than their untreated counterparts; moreover,
PCA concentration was unchanged (Fig. 2B).
P. aeruginosa
eDNA originates from the genomic DNA of
dead cells and is therefore high molecular weight and may be bound by other biomolecules (e.g.
proteins) (Kavanaugh et al., 2019). Therefore, DNase treatment was likely only partially effective
because it could not cleave ds DNA in the presence of other bound matrix components, and/or because i
t
did not have enough activity to degrade the eDNA completely to eliminate phenazine binding sites. We
also compared the phenazine retention in the WT to a Pel mutant
(
D
pel
) and found that biofilms without
Pel
retained significantly more PYO (Fig. 2C). Because Pel is known to bind eDNA (Jennings et al.,
2015), these results suggest that Pel may partially block access to eDNA by PYO, although this remains
to be tested
in vitro
.
To probe the eDNA binding sites within the biofilm using a different approach, we competed phenazines
against ethidium bromide, a classical DNA intercalator. Since PCN and PYO compete for DNA binding
sites with ethidium
in vitro
, and ethidium is largely excluded from cells (Jernaes and Steen, 1994)
, we
reasoned that these intercalators could compete for binding sites in the biofilm eDNA. We grew
∆
phz*
biofilms with 50
μ
M PCN and PYO and increasing amounts of ethidium in the underlying agar. Figure
2D shows that increasing concentrations of ethidium resulted in successively less PYO accumulating in
the biofilms, while PCN accumulated to a similar lower level in the presence of any amount of added
ethidium.
Confocal microscopy of
WT
and
∆
phz*
colony biofilms with
a c
ell -impermeable ds DNA dye, TOTO-1
(Okshevsky and Meyer, 2014), showed abundant eDNA localized in dead cells and in between cells
(Fig. S3D). We quantified the bulk concentration of eDNA in colony biofilms by incubating biofilm
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suspensions with TOTO-1 and measuring dye fluorescence. Both WT and
∆
phz*
biofilm suspensions
yielded large fluorescence values when incubated with TOTO-1. These values fall within the range of 60
– 500
μ
M bp ds DNA in the colony biofilms, when calibrated against standards of calf thymus DNA
(Fig. 2E). However, adding calf thymus DNA to the biofilm suspensions did not yield the expected
increase in dye fluorescence (Fig. S3E), which suggests that the dye may be partially inhibited by
biofilm components. Therefore, this order of magnitude estimate of biofilm eDNA should be interpreted
as a lower bound on the true value. Given this estimate, the biofilm eDNA (>100
μ
M bp) is in excess of
PYO (~80
μ
M), but it may not be in excess of PCN (>300
μ
M). Due to its poor aqueous solubility, i
t is
probable that PCN crystallizes extracellularly at the observed biofilm concentrations, which could lead
to its measured retention (Hernandez et al., 2004)
. Together, our
in vivo
and
in vitro
results are
consistent with eDNA providing binding sites for oxidized PCN and oxidized PYO in the biofilm
extracellular matrix.
Constraints on phenazine electron transfer mechanisms
in vitro
and
in vivo
Given that phenazines are differentially bound and retained in biofilm eDNA, we next sought to
constrain how electron transfer might be achieved in this context
. Previous research has shown distinct
localization patterns for different phenazines within biofilms, with the lowest potential phenazines (
e.g.
PCA) in the interior, and the highest potential phenazine (
e.g.
PYO) at the oxic periphery (Bellin et al.,
2014, 2016)
. To test whether electron transfer could occur between these molecules in solution, we
mixed different oxidized and reduced phenazines under anoxic conditions and monitored the absorbance
spectra before and after mixing (Fig. 3A-B). Because PYO exhibited the largest changes in absorbance
upon reduction, we monitored different mixtures of PYO with PCA or PCN
at 690nm (unique PYO
absorbance maximum) starting one minute after mixing, at which point equilibrium had been achieved.
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Reactions proceeded as expected from the redox potentials of the phenazines, where PYO was almost
completely reduced by the lower potential PCA and PCN, but reduction of PCA and PCN by the higher
potential PYO was minimal (Fig. 3B
, Fig. S4A). In addition to establishing that electron transfer can
occur between different phenazines, given their similar structures, these results suggest that electron
transfer between like phenazines (
e.g.,
PYO
-PYO electron self-
exchange) can occur within a redox
gradient. Moreover, reactions between reduced PCA or PCN and oxidized PYO proceeded faster than
oxidation of any of these phenazines by molecular oxygen (Fig. 3C). We next wondered whether the
presence of eDNA would affect the extent of PYO reduction. PYO reduction by PCA or PCN proceeded
to completion in the presence of eDNA (Fig. 3B). Because PCA does not bind eDNA, this result
suggests electron transfer is occurring in solution between PCA and unbound PYO. For PCN and PYO
that both bind eDNA, it is also possible that electron transfer is achieved by their unbound counterparts
in solution. However, it has long been known that DNA can facilitate electron transfer between bound
redox molecules (Genereux and Barton, 2010), motivating us to test whether such a process could also
occur within our
P. aeruginosa
biofilms.
DNA facilitate
s charge transfer (DNA CT) through the
p
-stacked base pairs (Genereux and Barton,
2010), and recent studies have shown that DNA CT can occur over kilobase distances (Tse et al., 2019)
.
A previous study suggested that PYO might be able to transport electrons via DNA (Das et al., 2015)
,
but given the preliminary nature of these experiments, we decided to revisit these experiments more
rigorously. To
better test the ability of phenazines to carry out DNA CT, we covalently attached a
phenazine via a flexible linker to one DNA strand and then made DNA-modified gold electrodes with a
thiol linker on the complementary strand according to standard protocols (Kelley et al., 1997; Slinker et
al., 2010, 2011). Specifically, short ds DNA molecules (17 bp) were covalently linked to the gold
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surface to form a packed monolayer, and the distal 5' end of each duplex contained a covalently linked
PCN
, the phenazine derivative most amenable to synthesis (see Materials and Methods
) (Fig. 3D
). Thus,
we established a well-defined chemical system to test if a phenazine could participate in electron
transfer to the electrode through the ds DNA
.
Because the efficiency of DNA CT depends upon the integrity of the
p
-stacking of base pairs within the
DNA duplex (Genereux and Barton, 2010), we compared well-matched duplex DNA monolayers to
duplex DNAs containing a single base mismatch that stacks less efficiently
(Fig.
3D
). We
utilized
multiplexed DNA chips to facilitate replicate comparisons between well-matched and mismatched DNA
monolayers (Fig. S4B-D); measurements with a non-
intercalating control probe showed that these
different monolayers had very similar surface coverages (Fig. S4E-F
) (Slinker et al., 2010)
. Figure
3E
shows that the mismatched construct yielded diminished current in the phenazine redox peak, consistent
with the charge transfer being DNA-mediated; the presence of the intervening mismatch inhibits
DNA
CT
. This mismatch effect was consistent across replicate low density (32-54% decrease) and high
density (36-69% decrease) DNA monolayers (Fig. S4C and supp. text). These results displayed
mismatch attenuation similar
to
that observed for well characterized small molecules shown to stack
with the DNA duplex and carry out DNA-mediated CT (Slinker et al., 2011). Strikingly, in the presence
of oxygen, a classic voltammetric signal characteristic of electrocatalysis was obtained (Fig. 3F)
centered on the phenazine redox peak. Hence the DNA-tethered phenazine is able to accept electrons
from the electrode through the DNA
p
-stack and then reduce oxygen in a catalytic fashion. Together,
these results demonstrate that a
P. aeruginosa
phenazine can participate in DNA CT
in vitro
.
To test if biofilm eDNA could support DNA CT, we incubated
∆
phz*
colony biofilms suspended in PBS
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with well-characterized intercalators that can perform DNA CT reactions, which can be monitored by
time
-resolved spectroscopy (Arkin et al., 1996a)
. The quenching of photoexcited Ru(phen)
2
dppz
2+
by
Rh(phi)
2
bpy
3+
is well-characterized and known to occur by a redox mechanism (Stemp et al., 1995) and
not energy transfer (e.g. FRET), with both the forward and back electron transfers occurring
predominantly on the picosecond timescale (Fig. 3G, Fig. S4G
) (Arkin et al., 1996b). Both of these
intercalators bind to ds DNA more than an order of magnitude more tightly than do phenazines.
Moreover, Ru(phen)
2
dppz
2+
is luminescent in aqueous solution only when intercalated in DNA (or
otherwise protected from water), precluding a 2
nd
order reaction between the complexes in solution.
Thus, in time-resolved emission experiments on the nanosecond
timescale,
static quenching, where
quenching
is fast
and occurs without a change in
the
Ru(phen)
2
dppz
2+
excited state
lifetime
, is consistent
with DNA-mediated CT (Arkin et al., 1996a), while slower dynamic quenching, which leads to a change
in emission lifetime, is consistent with a slower diffusive process
. We first compared the
Ru(phen)
2
dppz
2+
signals of liquid grown and biofilm suspensions (of the same optical density) to
determine if the signal was specific to eDNA; the ruthenium complex is not expected to be taken up by
the cells and bind to genomic DNA on the time scale of this experiment (Fig. 3H). We only observed
ruthenium luminescence in the presence of the biofilm suspension, consistent with ruthenium
luminescence being associated with binding to eDNA. We then examined the pattern of quenching of
Ru(phen)
2
dppz
2+
by Rh(phi)
2
bpy
3+
. Fig
ure 3I shows static quenching where the intensity of the
Ru(phen)
2
dppz
2+
signal decreases, while the observed decay kinetics are unchanged (Fig. S4H).
Therefore, we conclude that biofilm eDNA can support rapid DNA CT between these two metal
complexes, faster than the timescale for diffusion.
Electrode-grown biofilms retain PYO capable of extracellular electron transfer
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Having explored phenazine retention and electron transfer separately, we next wanted to establish a
system in which we could monitor both of these processes simultaneously
in vivo
. We took an
electrochemical approach to achieve this, growing
P. aeruginosa
biofilms on interdigitated
microelectrode electrode arrays (IDA) (Fig. 4A). Biofilms were grown by incubating IDAs in
bioelectrochemical reactors with planktonic cultures under oxic conditions and stirring (Fig. 4B). After 4
days, with medium replaced daily, mature biofilms were transferred to anoxic reactors with fresh
medium for electrochemical measurements. Confocal and SEM imaging revealed that the IDA biofilms
were heterogeneous in composition, but consistently contained multicellular structures of live cells and
abundant eDNA (Fig. 4C-
E,
Fig. S5A-B
). Under these conditions, the cells predominately produced
PYO, as measured by LC-
MS (
Fig. S5C). Because PYO was the most tightly retained phenazine in
colony biofilms and
in vitro
, we focused on this phenazine for the remainder of these experiments.
Originally used to measure conductivity of abiotic materials, the IDA has
a 2
-working electrode
geometry and recently
was
adapted to study EET through microbial biofilms (Boyd et al., 2015; Snider
et al., 2012; Xu et al., 2018; Yates et al., 2015)
. Measurements are made by driving electron transfer
between the two electrode bands across a 5
μ
m gap (Fig. 4F), which treats the biofilm like an abiotic
material, resulting in EET that is decoupled from the cells’ metabolic activity. Specifically, we used a
generator-collector (GC) strategy to measure EET through the biofilm, where the “generator” electrode
is swept from an oxidizing potential (E = 0mV vs. Ag/AgCl) to a reducing potential (E = -500mV),
while the “collector” electrode maintains a fixed oxidizing potential (E = 0mV). In this GC arrangement,
electron transfer into the biofilm from the generator occurs as the potential of the generator is swept
negatively, reducing PYO
(E
0
= -250mV) at the biofilm/generator interface. Electrons are conducted
across the gap through the biofilm due to EET, either by physical diffusion of PYO or other
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mechanisms. The electron transfer at both the generator and collector is measured as current (I
gc
) and
plotted against the potential of the generator electrode.
Implicit in generating a sustained I
gc
is recycling
of the redox molecules that support the observed current.
Figure 4G shows that WT and
D
phz
* + PYO
biofilms supported current (above background) across the 5
μ
m gap over a ~3 min scan as the generator
potential approached PYO’s redox potential (-250mV vs. Ag/AgCl), consistent with PYO-mediated
EET; significantly,
D
phz
* alone did not. Dependency of current on the generator potential observed here
is consistent with PYO-mediated EET being a diffusive process. Reduction of PYO at the generator and
oxidation of PYO at the collector generates a redox gradient that drives EET through the intervening
biofilm, resulting in current centered on the PYO midpoint potential that saturates at strongly reducing
generator potentials (Snider et al., 2012)
. Here the diffusive nature of EET would reflect either physical
diffusion of PYO through the biofilm (reduced
PYO
from the generator to collector, oxidized
PYO
from
the collector to the generator) or the effective diffusion of electrons through bound PYO (Strycharz-
Glaven et al., 2011a).
To test whether these short-term GC measurements of biofilm EET indicate long-term ability to support
metabolic activity, we poised the IDA electrodes at an oxidizing potential (+100mV vs. Ag/AgCl) and
monitored the current produced by the biofilm over 4 days in the presence of 40 mM succinate as the
electron donor for cellular metabolism under anoxic conditions. In this configuration the observed
current would be due to cellular oxidation of succinate coupled with PYO-catalyzed EET to the poised
electrode, where the electrode acts as the terminal electron acceptor instead of oxygen (Fig. 4H). Figure
4I shows that both WT biofilms relying solely on endogenous PYO and
D
phz
* biofilms + exogenous
PYO generate robust current over many days, while
D
phz
* alone does not. The daily periodic rise in
current likely reflects the impact of slight temperature or light fluctuations in the room on the cells’
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metabolic activity over the course of the experiment (Kahl et al., 2016, 2019). Differences in current
magnitude between the WT and
D
phz
* + PYO runs are likely due to differences in PYO abundance in
the biofilms and/or efficiency of cellular PYO reduction. Together, these results demonstrate that the
IDA biofilms can use retained
PYO
for EET to support metabolic activity.
To determine whether our IDA biofilms retained phenazines in the same manner as colony biofilms,
D
phz
* IDA biofilms were soaked in PYO in one reactor and then transferred to another reactor with
fresh medium lacking PYO (Fig. 5A). The equilibration of PYO (from the IDA biofilm) with the fresh
medium was monitored by square wave voltammetry (SWV) (Fig. 5B), for which peak current (I
swv
) is
proportional to the concentration of the PYO remaining in the biofilm at each time interval (Bard et al.,
1980). Thus, the rate of decay of I
swv
provides a means to assess the loss rate of PYO from the biofilm
, a
measure of how tightly PYO is retained. Upon transfer, the biofilm PYO peak current (I
swv
) decays in
30-45
min
, while for a blank IDA (no biofilm) dipped in PYO, I
swv
decays in 2-3 min (Fig. 5E). We
compared PYO and
PCA
soaks and found that PCA immediately became undetectable by SWV or GC
in the transfer reactor (
Fig. S5D-E) because it quickly diffuses out of the biofilm, as expected because it
does not bind eDNA. Thus, like colony biofilms, IDA biofilms appear to tightly retain PYO but not
PCA.
Electron transfer through biofilms is faster than PYO loss
Next, we sought to understand the efficiency of PYO-mediated EET in the biofilm. We reasoned that a
determination of “efficiency” would compare the rate of electron transfer (which supports the
metabolism of the O
2
limited cells) to the loss rate of PYO from the biofilm (which limits the utility of
each PYO molecule). These two processes can both be described by diffusion coefficients, so in a single
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electrochemical experiment we measured the apparent diffusion coefficient for the PYO-mediated EET,
D
ap
, which characterizes all of the redox processes with the electrode, and the diffusion coefficient for
PYO as it is lost from the biofilm, D
loss
.
We determined D
ap
for EET by PYO in an IDA
D
phz
* biofilm to avoid confounding PYO retention with
production, although WT biofilms with endogenous PYO yielded similar results (
Fig. S6A-
B).
Each
D
phz
* biofilm was soaked with PYO in one reactor then transferred to a second reactor lacking PYO
(Fig. 5A), and the equilibration of the biofilm PYO into the fresh medium was monitored by paired
SWV and GC measurements over time (Fig. 5B and Fig. 5C). Noting that I
swv
is proportional to
퐶
∗
#
퐷
%&
, where C is the effective PYO concentration (Bard et al., 1980), whereas I
gc
is proportional to
퐶
∗
퐷
%&
(Strycharz-Glaven et al., 2011b), plotting I
gc
vs. I
swv
for each time point is expected to yield a
linear dependency with a slope (
m
) proportional to
#
퐷
%&
(Fig. 5D) when D
ap
is independent of the
concentration of PYO in the biofilm (Akhoury et al., 2013; White et al., 1982a). In this manner, it
is
possible to measure D
ap
for PYO remaining in the biofilm at any given instance when its concentration
is unknown.
Applying this approach, for two biological replicates of
D
phz
* biofilms (three technical replicates each),
we found that the mean D
ap
for PYO is 6.4 x10
-6
cm
2
/sec
over a wide range of PYO biofilm
concentrations (Fig. 5F, Fig. S6C
). To validate our approach, we measured D
ap
using a blank IDA with
no biofilm, only solution PYO for which we expect D
ap
»
D
loss
. Using known PYO concentrations with a
blank IDA (no biofilm) we obtained nearly identical estimates (D
ap
»
6.8x10
-6
cm
2
/sec
) using our I
gc
vs.
I
swv
method (concentration unknown) or established methods (I
gc
vs. [PYO] and I
swv
vs. [PYO]) that
depend on known concentrations (Fig. 5F, Supplementary text, Fig. S6D-G) (Bard et al., 1980)
. We
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further validated our measurement scheme by comparing it to an established chronocoulometry
technique with two other redox molecules (Hexaammineruthenium(III) and ferrocenemethanol
) in the
presence and absence of the polymer Nafion. In all cases estimates of D
ap
by the two methods were
within ~2x (Fig. S6H).
To
estimate the upper limit for D
loss
of PYO lost from
P. aeruginosa
biofilms, we applied a semi-infinite
one-dimensional diffusion model (Supplemental Text) to fit the decay of the same I
swv
measurements
used above (Fig. 5B, 5E). Although each SWV scan depends on D
ap
, the decay
process of I
swv
results
from loss of PYO out of the biofilm
(D
loss
). This calculation requires a scan time constant, whose value
we constrain by using the blank control where we assume D
ap
= D
loss
, allowing us to solve for this
constant (see Supplemental Text and Fig. S7A).
For
D
phz
* biofilms, this diffusion model yields a
mean
D
loss
of 2.0x10
-7
cm
2
/sec (
Fig. S7B). Hence, D
ap
for PYO
-mediated biofilm EET is more than 25-fold
higher than D
loss
(Fig. 5F). While this model simplifies the actual physical system, it provides a means to
estimate an upper limit for the rate of PYO loss from the biofilm
. Such a low D
loss
is consistent with the
relatively long time it takes for I
swv
to decay when transferred to fresh medium for PYO in a biofilm
(~45min) compared to PYO for a blank IDA (<2min) (Fig. 5E) or to PCA in a biofilm (< 1 min) (Fig.
S5E
). Moreover, the conservative assumptions of our model make it likely that we have underestimated
the true difference between D
loss
and D
ap
in the biofilm.
Collectively, these observations support the idea
that PYO EET occurs rapidly compared to the loss of PYO diffusing out of the IDA biofilms.
DISCUSSION:
The redox activity of phenazine metabolites produced by
P. aeruginosa
has intrigued researchers since
the 1930’s (Friedheim, 1934), and over the last twenty years a model has emerged that links phenazine
electron transfer to biofilm metabolism (Dietrich et al., 2013b; Hernandez and Newman, 2001; Jo et al.,
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2017). Using the well-characterized
P. aeruginosa
-phenazine system as a model for studying EET
mediated by extracellular electron shuttles, here we addressed the conundrum of how phenazines
complete their redox cycle within the biofilm matrix without being lost to the environment. Our results
point to eDNA as being a critical component of the matrix that facilitates phenazine cycling for EET.
Quantifying phenazine retention in a simple biofilm system was our first goal. While past work has
measured phenazines in culture supernatants and in agar underlying colony biofilms (Bellin et al., 2014,
2016; Dietrich et al., 2006), phenazine concentrations within any type of biofilm system were unknown.
We found that the degree of retention varied dramatically between the three studied phenazines in
colony biofilms, with PCN and PYO being strongly enriched in the biofilm, in contrast to PCA, which
readily diffuses away. We observed a similar trend for biofilms grown in liquid medium attaching to
IDAs. No
tably, oxidized PCN and PYO bind ds DNA
in vitro
, and perturbing extracellular DNA
binding sites disrupted PYO retention (and PCN to a lesser extent)
in vivo
. Previous studies have shown
PYO actually promotes eDNA release via cell lysis, so PYO retention by eDNA suggests a connection
between eDNA production and utilization. To our knowledge, PYO retention in eDNA is the first
example of a metabolically helpful (as opposed to biofilm structural) molecule being bound by the
extracellular matrix of a biofilm. Because eDNA is found in biofilms made by diverse species and many
small molecules exhibit DNA binding capacity, our results may be broadly relevant to diverse biofilm
functions involving extracellular metabolites.
Recognizing that a primary function for phenazines is extracellular electron transfer, we characterized
this process
in vivo
. Our IDA experiments suggest that PYO simultaneously can be retained (low D
loss
)
and facilitate fast EET (high D
ap
), establishing the efficiency of this process. To understand the
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mechanism that underpins this efficient EET, we can interpret our results in the context of electron
transfer theory from the study of redox polymers (Dalton and Murray, 1990). This theory holds that the
measured D
ap
is the sum of the effective diffusion coefficient of electrons due to
self
-exchange reactions
among bound shuttles
(D
e
) and the physical diffusion coefficient
(D
phys
) of any unbound shuttles (D
ap
=
D
e
+ D
phys
). In this context, we think there are two ways to explain our electrochemical results that
biofilm PYO D
ap
(~6.4 x10
-6
cm
2
/sec
) is higher than biofilm PYO D
loss
(~2.0
x10
-7
cm
2
/sec
), and similar
to solution PYO D
phys
( 6.8 x10
-6
cm
2
/sec
) (Fig. 5F)
. First,
if we
assume our measured D
loss
is the same as
PYO D
phys
within the biofilm, the difference between D
ap
and D
phys
can be explained by D
e
. Our
in vitro
data suggest that such
self
-exchange reactions (D
e
) could be DNA-mediated. In the case of very rapid
electron transfer among eDNA-bound PYO, D
ap
would still be limited by counter ion diffusion,
consistent with a measured D
ap
similar to that of a small molecule in solution (~7 x 10
-6
cm
2
/s)
(White et
al., 1982b)
. Alternatively, the measured loss of PYO from the biofilm, D
loss
, may not accurately reflect
physical diffusion of PYO within the biofilm, D
phys
. Because the IDA biofilms do not completely cover
the electrodes, PYO may be able to physically diffuse in solution adjacent to them. In this case, diffusion
in solution would be consistent with the measured D
ap
, and the low D
loss
measurement would reflect the
slow release of PYO from the IDA biofilm due to its retention by eDNA. Regardless, the striking
measured difference between D
ap
and D
loss
indicates that PYO electron transfer promotes efficient
biofilm EET metabolism because electron transfer occurs rapidly, while loss of PYO to the environment
is slow.
Together, our results allow us to refine our model for how phenazine EET may operate within biofilms
(Fig. 6). Intriguingly, PYO biosynthesis requires O
2
, whereas PCN and PCA do not, and electrochemical
imaging underneath colony biofilms has shown lower potential phenazines in the anoxic interior (PCA,
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PCN) and the higher potential phenazine (PYO) near the oxic periphery (Bellin et al., 2014, 2016)
.
Reduced PYO is also known to react with oxygen significantly faster than PCA and PCN (Wang and
Newman, 2008). As such, it is tempting to speculate that phenazines are ordered in the biofilm matrix in
a sequence akin to a large extracellular electron transport chain—from reduced PCA/PCN in the anoxic
interior, to eDNA bound PYO at the oxic periphery, and ultimately to molecular oxygen (Fig. 6A). How
then do phenazines exchange electrons within this framework? Noting that the heterogeneity of the
biofilm matrix makes it possible that different electron transfer pathways could occur in these complex
systems, we favor two mechanisms mediated by eDNA for how phenazine EET may operate in the
matrix (Fig. 6B). Both mechanisms assume that reduced PCA and PCN will diffuse from the anoxic
zone to the oxic zone
(i), where PYO
ox
is
bound to eDNA. In one model (Fig. 6B top), PYO’s binding
equilibrium will result in some PYO
ox
unbinding from the eDNA, allowing it to
react with
PCA
red
or
PCN
red
(ii).
PYO
red
then
react
s with oxygen generating
PYO
ox
(iii), which rebinds
DNA.
In the other
model (Fig. 6B bottom), reduced phenazines (likely PCN) intercalate into eDNA and reduce PYO
ox
via
DNA CT
(ii). PYO
red
unbinds DNA, reacts with oxygen
(iii),
and PYO
ox
rebinds DNA. Given that
PCA
red
and PCN
red
react more quickly with PYO
ox
than O
2
, then both models would facilitate the re-
oxidation of the interior phenazines and promote diffusion back toward the anoxic interior (iv). These
non-mutually exclusive models integrate what is known about phenazine O
2
reactivities, redox
potentials and biosynthesis zones, and help explain how PYO and eDNA interactions enhance EET in
P.
aeruginosa
biofilms by facilitating retention and electron transfer
. Testing these models in
physicochemically well-defined matrixes in addition to complex biofilms represents an exciting
challenge for future research.
In conclusion, our findings illustrate that
eDNA
binding provides a mechanism to resolve how otherwise
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diffusible extracellular electron shuttles can catalyze efficient EET in real world, open systems. Beyond
serving as a structural support, carbon source, or genetic reservoir, our studies reveal that interactions
between extracellular electron shuttles and eDNA in biofilms underpin their metabolic vitality. It
is
noteworthy that eDNA is abundant in many biofilms (Flemming and Wingender, 2010) and diverse
biofilm-forming bacteria have the potential to produce
extracellular electron shuttles
(Glasser et al.,
2017a). Accordingly, eDNA retention of these electron shuttles—and perhaps of other biologically
useful molecules—may
represent a widespread strategy whereby a reactive extracellular matrix supports
bacterial biofilms in unexpected and physiologically significant ways.
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Ack
nowledgments:
We thank Jeanyoung Jo, Lars Dietrich and Matthew Parsek for providing strains.
This work was supported by grants to D.K.N. from NIH (1R01AI127850- 01A1) and ARO (W911NF-
17-1-0024), to J.K.B. from NIH (GM126904), and to S.H.S., D.K.N and J.K.B. from the Rosen
Bioengineering Center at Caltech. E.C.M.T. was supported by a Croucher Foundation Research
Fellowship.
Author contributions:
Conceptualization, S.H.S., J.K.B., L.M.T., and D.K.N.;
Methodology, S.H.S., E.C.M.T., M.D.Y., F.J.O., S.A.T., E.D.A.S., J.K.B., L.M.T., D.K.N.; Formal
Analysis, S.H.S. and L.M.T.; Investigation, S.H.S., E.C.M.T., M.D.Y., F.J.O., S.A.T., E.D.A.S.;
Resources, J.K.B., L.M.T. and D.K.N.; Writing – Original Draft, S.H.S. and D.K.N.; Writing –
Reviewing & Editing, S.H.S., E
.C.M.T., M.D.Y., F.J.O., S.A.T., E.D.A.S., J.K.B., L.M.T., D.K.N.;
Visualization, S.H.S.; Supervision, J.K.B., L.M.T. and D.K.N.; Funding Acquisition, J.K.B., L.M.T. and
D.K.N.
Competing interests:
Authors declare no competing interests
.
Data and materials
availability:
Data and strains used in this study are available on request. Data and code are available at
github.com/DKN-lab/phz_eDNA_2019.
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