DESIGN AND CHARACTERIZATION OF CYANOBACTERIAL BIOREPORTERS TO MEASURE PHOSPHORUS AVAILABILITY IN MARINE SYSTEMS

Mark Jeremy Rozmarynowycz

A Thesis

Submitted to the Graduate College of Bowling Green State University in partial fulfillment of the requirements for the degree of

MASTER OF SCIENCE

December 2009

Committee:

R. Michael L. McKay, Advisor

George Bullerjahn

Scott Orland Rogers

ii

ABSTRACT

Dr. Robert Michael McKay, Advisor

Picocyanobacteria are major primary producers in pelagic waters, and therefore

an excellent indicator of the productivity of an environment. This highlights their functionality in identifying and monitoring factors constraining such productivity. While

inorganic P can be easily tested with chemical tests, the potential use of organic P is

neglected. By constructing whole cell bioreporters relevant to the open ocean, the

environment can be monitored from the perspective of a microorganism. In this study

two new bioreporters were constructed using the pelagic strain Synechococcus spp.

WH8102. Two genes sensitive to early P-stress were chosen for fusion of their promoter

regions to a luxAB cassette. The reporter strain OPD contains the promoter sequence of phoD, an alkaline phosphatase, while the reporter strain OPN contains the promoter

sequence of nucA, a 5 nucleotidase. These strains were created by constructing new

plasmid vectors, pOPD and pOPN, capable of conjugation into Synechococcus spp.

WH8102. Initial characterization shows a significant difference (P < 0.05, t-test)

between luminescence of a strain grown in P deplete medium (200 nM) vs. P replete

medium (80 μM) after 2 days of incubation. After the addition of 80 μM P, luminescence

levels dropped to levels similar to those grown in P replete conditions. Initial

characterization of this strain demonstrates its potential use in detecting bioavailable P, as

well as providing a real time view of the environment from a microorganism’s

perspective. iii

Dedicated to my parents,

Mary and Walter Rozmarynowycz. iv

ACKNOWLEDGMENTS

I would like to thank my advisor, Dr R. Michael L. McKay, and the members of my committee, Dr George S. Bullerjahn and Dr Scott Orland Rogers. This thesis would not be possible without your advice and guidance.

I would also like to thank the members of the Bullerjahn and McKay labs: Irina

Ilikchyan, Mike Schlais, Olga Kutovaya, Alex Nazarov, Maitreyee Mukherjee, Nigel

Dsouza, Korinna Straube, and Brian Schmidt. Thank you for your help and support, your advice was invaluable.

I would also like to thank to my professors at Lorain County Community College, especially Dr. Kathryn Durham who got me involved in bioreporter research as an undergraduate. I am also grateful to Dr. Harry Kestler and Dr. Adam Miller for their support and contributions to my undergraduate research.

I would also like to thank my family, especially my wife Clair, for their support and patience.

This material is based upon work supported by the National Science Foundation

under Grant No. 0727644

v

TABLE OF CONTENTS

Page

INTRODUCTION ...... 1

Considerations for bioreporter construction ...... 2

Reporter genes...... 3

Construction strategies...... 6

Genetic transformation strategies: prototype reporter strains...... 6

Conjugative strategies for marine Synechococcus spp...... 8

Cyanobacterial bioreporters as environmental sensors ...... 11

Bioreporters of trace metal availability: Fe ...... 11

Bioreporters of N availability...... 14

Bioreporters of P availability ...... 17

Issues and challenges in assessing bioavailable P...... 17

P-Responsive bioreporters ...... 20

METHODS AND MATERIALS...... 22

Media and growth conditions...... 22

Promoters……...... 22

Plasmid construction...... 23

Conjugation into Synechococcus...... 24

Bioreporter Characterization...... 27

RESULTS………………...... 28 vi

Verification of plasmid insertion...... 28

Growth of wild type vs. bioreporter strains ...... 28

Delivery of aldehyde substrate to the bioreporter ...... 29

P-Responsive luminescence of bioreporter strains...... 32

DISCUSSION………………...... 34

CONCLUSION………………...... 38

REFERENCES ...... 40 vii

LIST OF FIGURES/TABLES

Figure Page

1 Map of pALPHA, pRL153, and pOPD ...... 26

2 Colony PCR verification of plasmid insertion in Synechococcus spp. WH8102 ..... 30

3 Growth of bioreporter strains OPD and OPN, compared with the growth of wild type

Synechococcus spp. WH8102 ...... 31

4 Comparison of solvent type and concentration of n-decyl aldehyde concentrations on cell

luminescence...... 33

5 Luminescence expression plotted with growth curves for nucA and phoD bioreporters in

both P sufficient and -depleted conditions...... 35

Table Page

1 Primers used in this study ...... 25

2 Plasmids used in this study ...... 25

1

Introduction

There is no photic environment on Earth in which the presence of cyanobacteria has not been documented (Whitton and Potts 2000). From pole to pole, cyanobacteria can be major primary producers in both aquatic and terrestrial ecosystems. This is particularly true in oligotrophic marine and freshwater systems, where picocyanobacteria dominate the pelagic biomass (reviews, Ting et al. 2002, Callieri and Stockner 2002). Indeed, it is estimated that in the open sea, the picocyanobacteria Prochlorococcus spp. and Synechococcus spp. are responsible for up to 80% of the primary production (Goericke et al. 1993, Liu et al. 1997,

Veldhuis et al. 1997). Similar values have been documented for oligotrophic large lakes

(Fahnenstiel and Carrick 1992, McKay et al. 2005a, Ivanikova et al. 2007b). Given their ubiquity and position at the base of the food web, it comes as no surprise that cyanobacteria are viewed as excellent proxies for the productivity of the aquatic environment (e.g. Silvert, 1996, Ware and

Thompson, 2005). Such a fundamental position in the functioning of an ecosystem makes cyanobacteria ideal targets for identifying and monitoring factors that may constrain such productivity. As a consequence, genetically engineered cyanobacterial bioreporters have been developed for this purpose.

Specifically, a bioreporter is a cell designed to produce a quantifiable signal in response to a specific change in the environment of that cell (Bachmann 2003, Belkin 2003). A regulated promoter element is fused to a gene whose product will yield a readout activity that can be monitored in real time. Typically, the readout is detected as luminescence from a luciferase activity (Andersson et al. 2000, Mackey et al. 2007), or fluorescence of a GFP derivative (Kunert et al. 2000). Thus, environmental conditions can be examined from the perspective of a living cell, complementing parallel chemical and physical measurements of the environment in 2

question. Overall, bioreporters provide information on the bioavailability or toxicity of a

specific analyte to the cell, information that chemical measures of concentration and speciation

cannot provide. In our laboratory and others, efforts have focused on engineering picocyanobacteria designed to report on the bioavailability of macro- and micronutrients, specifically phosphorus (P), combined nitrogen (N) and iron (Fe) (Schreiter et al. 2001, Durham et al. 2002, Gillor et al. 2002, Mbeunkui et al. 2002, Durham et al. 2003, Gillor et al. 2003,

Ivanikova et al. 2005, Boyanapalli et al. 2007). Bioreporter construction and characterization has become increasingly straightforward, as there are a number of cyanobacterial strains amenable to genetic manipulation, and the wealth of genomic information has enabled the rapid identification and subcloning of specific target promoter sequences responsive to environmental stimuli. In this chapter, we will discuss the construction and use of cyanobacterial bioreporters, taking into consideration the reporter system, the strain to be employed and the assay methods necessary to yield reliable data on the specific parameter to be measured.

Considerations for bioreporter construction

Characteristics of bioreporters vary depending on the experimental conditions and detection methods employed. The choice of reporter genes depends on factors such as expression efficiency, stability, background activity, detection method and assay methodology (Daunert et al., 2000). Advantages of using bioreporter technology in environmental monitoring are sensitivity, selectivity and stability, all of which provide ideal conditions to assay environmental samples. Another important advantage is their ability to provide physiological information pertaining to the whole cell that is not possible with methods involving cell extracts. 3

Implementing the use of bioreporters can be a cost-effective, simple, and environmentally benign

process, which facilitates on-site monitoring (van der Meer et al. 2004). Limitations of

bioreporters include concerns pertaining to their ecological relevance to a given system as well

as situations where the analyzed compound may interact with extracellular material and alter the chemistry of an environmental sample (Harms et al. 2006). Therefore, when choosing a bacterial reporter strain, it is vital to consider various environmental factors to design an informative reporter strain. In this section we will address the many variables to be encountered while developing a cyanobacterial bioreporter.

Reporter genes

An initial consideration is the type of reporter gene to be employed. Whereas the majority of reporter fusions to date have been constructed with the Vibrio harveyi luxAB luciferase genes, by no means is that the only option. For example, GFP-based reporters have been developed in filamentous cyanobacteria to examine temporal gene regulation during heterocyst development

(review, Golden and Yoon 2003), and platforms to yield GFP fusions for Synechocystis sp. strain

PCC 6803 (Kunert et al. 2000) and Prochlorococcus spp. (Tolonen et al. 2006) have been developed in the past decade. Additionally, eukaryotic luciferase genes (luc, pxvGR, pxvRE)

have been employed to monitor circadian gene expression (Kitayama et al. 2004, Mackey et al.

2007). Given the fact that bacterial and the varied eukaryotic luciferases have distinct chemistry,

substrate requirements and emission wavelengths, the characteristics of these enzymes allows for

the development of multichannel bioreporters in which readouts from several promoters can be

recorded from the same cell. 4

Each fusion gene and gene product has particular advantages, depending on the application. GFP and other fluorescent proteins require no substrate or redox chemistry, thus the accumulation of the folded protein yields a direct readout signal. GFP and YFP derivatives typically emit at wavelengths (500-540 nm) that do not overlap with absorption spectra of photosynthetic pigments. However, in aquatic systems, fluorescent cyanobacterial bioreporters are of limited utility when fluorescent organic matter is present that can yield a high signal-to- noise ratio (N.V. Ivanikova and G.S. Bullerjahn, unpublished data). This is of particular concern in fresh waters and coastal marine environments, where fluorescent CDOM (emission 400 – 550 nm) can comprise a significant fraction of total DOC (Coble et al. 1990, De Souza Sierra et al.

1994, Boyd and Osburn, 2004, Kowalczuk et al. 2005). By contrast, measurement of luminescence from luciferase fusions is not affected by interfering organic compounds, at least not directly (see Hassler et al. 2009 for discussion). Additionally, studies with gfp and dsred fusions in Escherichia coli have shown that bioreporter response is slower than luciferase constructs, possibly limited by the slow rate at which fluorescent proteins fold into their functional conformation (Hakkila et al. 2002). In light of these issues, our use of gfp has been restricted to fusion constructs to a constitutive promoter (K.K. Brinkman and G.S. Bullerjahn, unpublished). In concert with a regulated luciferase reporter fusion, the population size of the reporter strain in an environmental sample can be monitored by GFP fluorescence in a flow cytometer or by epifluorescence microscopy.

Luciferase fusions, both luxAB and luc, involve the expression of a monooxygenase that requires an organic substrate and molecular oxygen. Even though the chemistry of the two classes of enzymes are vastly different, the nature of luciferase fusions requires that the bioreporter cells are metabolically active and in an aerobic environment. Availability of the 5

appropriate substrate is also an important consideration. In the case of bacterial luciferase, the

luxCDE genes direct the synthesis and regeneration of the acyl aldehyde substrate. In the pioneering work by Susan Golden’s lab, bacterial luciferase fusions were generated to explore

Synechococcus elongatus PCC 7942 circadian gene expression. In such strains, the Vibrio sp.

luxCDE genes are under constitutive control, allowing endogenous substrate for long-term real-

time monitoring of luminescence (Liu et al. 1995). However, such constructs often do not

produce saturating concentrations of substrate, requiring addition of exogenous decyl aldehyde to maximize luminescence (Durham et al. 2002, Porta et al. 2003). To date, cyanobacterial luc,

pxvGR and pxvRE fusions bearing the eukaryotic luciferase genes have relied on exogenous

luciferin added to the assay medium (Shao et al. 2002, Kitayama et al. 2004, Mackey et al. 2007)

as well as an energy requirement in the form of ATP. Bacterial luciferase emits at 490 nm

(Hastings 1978), and eukaryotic luciferases PxvGR, Luc and PxvRE emit in the green (549 nm,

560 nm) and red (622 nm), respectively, allowing spectral discrimination among different

promoter fusions in the same cell (Kitayama et al. 2004, Mackey et al. 2007).

Whereas luc and other eukaryotic luciferases are important tools in the construction of

multichannel cyanobacterial bioreporters, our lab has traditionally relied on the construction of

luxAB promoter fusions. Such strains yield a robust luminescent signal detectable from

bioreporter densities mimicking that of pelagic picocyanobacterial populations (105 mL-1 or ca.

1
g chl a mL-1). Below we describe the construction and use of freshwater and marine

bioreporters to assess aquatic nutrient bioavailability.

6

Construction strategies

Depending on the regulation of the promoter to be employed in bioreporter construction,

luciferase-dependent reporters can be designed to respond specifically to an analyte yielding

either an increase in luminescence (Class I, or “Lights on”), or a decrease in light emission

(Class II, or “Lights off”) (Belkin 2003, van der Meer et al. 2004). In our laboratory, we have

constructed both types and successfully used them to assess Fe (Class II; Durham et al. 2002,

2003, Boyanapalli et al. 2007) and nitrate bioavailability (Class I; Ivanikova et al. 2005). In this

section we discuss strategies for bioreporter construction, taking into consideration the host

strain and gene transfer protocols.

Genetic transformation strategies: prototype reporter strains

The first cyanobacterial reporter strains constructed used the freshwater model strains

Synechococcus elongatus PCC 7942 (Durham et al. 2002, Gillor et al. 2002, Durham et al. 2003,

Gillor et al. 2003) and Synechococystis sp. PCC 6803 (Ivanikova et al. 2005) and the halotolerant

Synechococcus sp. PCC 7002 (Boyanapalli et al. 2007). To date, bioreporter strains developed at

Bowling Green State University and used with success include Fe-dependent Class II

Synechococcus spp. PCC 7942 and PCC 7002 bioreporters that utilize the Fur-regulated isiAB

promoter (Durham et al. 2002, Boyanapalli et al. 2007), and the irpA promoter, part of the Fe-

responsive IdiB regulon (Yousef et al. 2003, Hassler et al. 2006, Nodop et al. 2008). Also

developed has been a Synechocystis sp. PCC 6803 Class I reporter employing the NtcA/NtcB-

regulated nirA (nitrite reductase) promoter (Ivanikova et al. 2005). Such strains have been employed to assess Fe (e.g. Boyanapalli et al. 2007; Hassler et al. 2009) and nitrate 7

bioavailability (Ivanikova et al. 2007a) in environmental samples. Since these strains can be

manipulated by efficient natural genetic transformation, promoter fusion plasmids can be

constructed and introduced into host strains as naked DNA. Furthermore, the complete genome

sequences available from these strains allow the amplification of any promoter for fusion

construction. Given the ease of this approach, and an existing genetic toolbox of luxAB promoter probe plasmids (Andersson et al. 2000, Kunert et al. 2000, Mackey et al. 2007), a suite of

Synechococcus PCC 7942 and Synechocystis sp. PCC 6803 bioreporters were rapidly developed

and characterized. Modification of the S. elongatus promoter fusion plasmid pAM1414

(Andersson et al. 2000, Mackey et al. 2007) yielded a suite of similar vectors for Synechococcus

sp. PCC 7002 (Boyanapalli et al. 2007). Despite the fact that these strains are not particularly

ecologically relevant with respect to most oligotrophic aquatic environments, these prototype strains afforded an opportunity to demonstrate the feasibility in using reporters in both freshwater (Durham et al. 2002, Porta et al. 2003, McKay et al. 2005a, Porta et al. 2005, Hassler et al. 2006, Ivanikova et al. 2007a, Hassler et al. 2008, Hassler et al. 2009) and marine samples

(Boyanapalli et al. 2007).

In general, the construction strategy involves the use of a plasmid vector incapable of replication in the cyanobacterial recipient strain. Chromosomal sequences engineered into the plasmid that flank a promoter::luxAB fusion and an antibiotic resistance marker allow the selection of drug-resistant survivors that have acquired a stable copy of the fusion construct

through homologous recombination (Liu et al. 1995, Andersson et al. 2000, Kunert et al. 2000,

Boyanapalli et al. 2007, Mackey et al. 2007). Often, the site for integration into the chromosome

is a so-called neutral site (NS), chosen because integration yields no observable phenotype

(Kunert et al. 2000, Mackey et al. 2007). Indeed, combining two or more NS integrative vectors 8

allow the construction of strains that express a promoter fused to luxAB at one site (NS1) and

luxCDE under constitutive expression at another (NS2). Alternatively, multiple luciferases can

be expressed by combining several NS insertions in the same strain (Mackey et al. 2007). By

contrast, recombination sites can be designed that yield a useful discernable phenotype. For

example, the Synechococcus sp. PCC 7002 promoter probe plasmids recombine into the desB

gene, encoding a fatty acid desaturase (Sakamoto et al. 1997, Boyanapalli et al. 2007). The

resulting construct is incapable of growth at temperatures below 15 °C, thereby reducing the

likelihood that the strain will grow if inadvertently released into the field.

Conjugative strategies for marine Synechococcus spp.

A long-term goal of our lab has been focused on the development of a suite of

picocyanobacterial reporters in strains that are ecologically relevant to the pelagic open sea.

Complicating this aim is the lack of a genetic transformation system for marine Synechococcus

spp. or Prochlorococcus spp. As a result, plasmid constructs are delivered into Synechococcus

spp. by conjugation from an E. coli donor (see below). Additionally, considerations must be

made regarding the type of strain to employ as a bioreporter host strain. Indeed, the availability

of several picocyanobacterial genome sequences yields a potentially difficult choice. Ideally, the best candidate would be a genetically manipulable strain whose genome has been sequenced, and for which nutrient assimilation pathways have been analyzed in silico (Palenik et al. 2003, Su et al. 2003). Meeting all these criteria is the strain Synechococcus sp. WH8102, a motile, pelagic

marine generalist (Palenik et al. 2003) and a member of the picocyanobacterial clade that

includes both marine Synechococcus spp. and Prochlorococcus spp. (Urbach et al. 1998). This 9

thesis contributes to our objective of developing bioreporter systems in this strain by mobilizing luxAB promoter fusions by conjugation.

As mentioned above, marine picocyanobacteria cannot be manipulated by genetic transformation. Instead, foreign DNA is introduced by conjugation from Escherichia coli donors bearing derivatives of broad host range plasmids capable of efficient transmission among divergent bacterial taxa (eg. Brahamsha 1996, Larsen et al. 2002). The E. coli donor strain contains a plasmid (e.g. pRK24) bearing the broad host range transfer functions required for conjugation. Indeed, efficient conjugational transfer of DNA has been documented for

Synechococcus sp. WH7803, WH8102 and WH8103 (3-6
10-3 per recipient; Brahamsha 1996) and Prochlorococcus sp. MIT9313 (Tolonen et al. 2006). For conjugation into these strains, broad host range plasmids derived from IncP (RP4 or RK2) and IncQ (RSF1010) incompatibility groups have been employed, thus these plasmids can be used as templates for the development of transmissible vectors bearing a lux promoter fusion capable of recombination into the chromosome. Alternatively, the promoter fusion can be delivered on a plasmid that can replicate in the recipient Synechococcus sp. strain at low copy number. Specifically, IncQ plasmid pRL153 has been used in matings with Synechococcus spp. (Brahamsha 1996) and

Prochlorococcus sp. MIT9313 (Tolonen et al. 2006) to yield an extrachromosomal promoter fusion plasmid. The availability of this plasmid system allows a rapid means for delivering and testing prototype promoter fusions into Synechococcus sp. WH8102 with high efficiency.

To yield a suicide plasmid capable of recombination with the chromosome, an RP4 or RK2 plasmid derivative maintained in the E. coli donor strain provides broad host range transfer functions allowing the delivery of a second plasmid into the cyanobacterial recipient. For our 10

purposes, E. coli BW20767 bearing plasmids pRK24 and pRL528 (Brahamsha 1996, McCarren and Brahamsha, 2005) is suitable to provide the transfer functions. To achieve this, a number of sequences must be present on the suicide plasmid. For conjugative transfer to WH8102, an appropriate oriT (RP4/R6K) compatible with the RP4 transfer functions provided by the E. coli donor strain (Larsen et al. 2002) is employed on the delivery suicide plasmid. The plasmid also contains a convenient cloning site adjacent to the promoterless luxAB genes and an antibiotic resistance marker, which are in turn flanked by chromosomal sequences allowing site-specific insertion into the chromosome by homologous recombination. Promoter fusion constructs are identified as spectinomycin resistant survivors arising following patch conjugation on SN plate media and subsequent selection on antibiotic-containing pour plates (McCarren and Brahamsha

2005).

Last, consideration must be given to the recombination site into the WH8102 genome. Ideally, the promoter-probe platform insertion should yield a neutral phenotype with respect to the physiological processes assayed by the bioreporter. One possibility is to use internal sequences of the swmB gene (ca. 800 bp flanking DNA) as a site for recombination. swmB is a large gene

(32 kb) essential for swimming motility in WH8102, a trait not widely distributed among the picocyanobacterial clade (McCarren and Brahamsha 2005). Due to the low GC content of swmB compared to the rest of the genome, it has been proposed that the gene was acquired by this strain via horizontal gene transfer (McCarren and Brahamsha 2005). swmB represents a candidate gene as a recombination target because interruption of the gene does not affect the strain’s capability to function as a bioreporter, yet the genetic stability of the chromosomal insertion can be assessed by examining the behavior of the construct in pour plates. A stable site-specific insertion yields small, non-motile colonies, phenotypically distinct from wild type. 11

Expanding bioreporter construction to other Synechococcus strains inhabiting distinctly different

marine environments is another long-term goal. For example, Synechococcus CC9311 is well

adapted to a coastal environment by bearing a large complement of genes encoding functions for

metal ion utilization and storage (Palenik et al. 2006). Conversely, fewer genes comprise the P

regulon encoding enzymes involved in phosphorus scavenging, reflecting the generally higher P

bioavailability in coastal regions. Since CC9311 is also amenable to genetic manipulation by conjugation, a suite of reporter constructs in this strain would provide additional insights into how coastal members of the picocyanobacterial clade sense nutrient fluxes compared to pelagic forms. This additional approach would also help determine the long-term validity of PCC 7002 strains that we have successfully manipulated as bioreporters for coastal environments.

Cyanobacterial bioreporters as environmental sensors

Bioreporters of trace metal availability: Fe

The role of Fe in physiological processes such as photosynthesis, respiration, and nitrogen assimilation makes it one of the most important nutritive factors for phytoplankton growth.

Despite its high abundance in the Earth’s crust, low Fe availability has been shown to limit phytoplankton growth in diverse marine environments (Hutchins et al. 1998, Boyd et al. 2007).

This apparent contradiction is attributed to features of Fe biogeochemistry that lead to precipitation or complexation of Fe species in oxic waters. These considerations, combined with regionally low aeolian input of Fe to many open ocean environments (Jickells et al. 2005) result in Fe deficiency to the endemic phytoplankton. 12

Efforts focused on studying Fe biogeochemistry over the past several decades have been biased

toward global surveys providing reliable measures of total dissolved (DFe) and particulate Fe

(PFe). Data collected show that DFe is present often in subnanomolar concentrations in open

ocean surface waters. Although absolute measures of DFe have been made in these regions, the

proportion of the total Fe pool that is available to the phototrophic plankton is poorly known.

Further, whereas chemical measures of total DFe can provide an estimate of the potential for Fe limitation, DFe need not be synonymous with bioavailable Fe. Some forms of PFe appear to be

bioavailable (Hassler et al. 2008, 2009) whereas some Fe associated with the operationally-

defined dissolved phase (< 0.45
m) is not immediately available for uptake (Hutchins et al.

1999). The complex chemical speciation of Fe in aquatic systems and the uncertainties

associated with biological assimilation of Fe species make it difficult to ascertain the fractions of

chemically detectable Fe that are readily available to phytoplankton.

Numerous approaches have been used to measure Fe availability in aquatic systems. Enrichment

bioassays, though offering direct experimental evidence for growth limitation, do not completely

mimic the undisturbed natural environment: grazing is disrupted, physical mixing is decreased

and the phytoplankton are isolated at a fixed optical depth (Carpenter 1996). To overcome these

concerns, several biochemical and molecular approaches have been suggested such as

monitoring the expression of Fe-responsive genes in environmental samples (Geiss et al. 2001,

2004), measuring the ratios of the redox catalysts ferredoxin:flavodoxin (Xia et al. 2004, McKay

et al. 2005b) or other suitable Fe-responsive proteins such as IdiA (Webb et al. 2001) and

assaying variable chlorophyll fluorescence (Behrenfeld et al. 2006). Implementing a living

system such as a bioreporter organism can help us gain a better understanding of the availability

of Fe from the perspective of a living cell. 13

A prototype Fe-responsive bioreporter was developed from the unicellular freshwater cyanobacterium S. elongatus PCC 7942 in a construct bearing a genetic fusion of the Fe- responsive isiAB promoter to the Vibrio harveyi luxAB genes encoding bacterial luciferase

(Durham et al. 2002). PisiAB is regulated in part by the Fe-dependent repressor Fur, yielding a

Class II (“Lights off”) reporter strain (Belkin, 2003). Characterization of this strain, KAS101, demonstrated the luminescent response to be a function of the free ferric ion concentration in metal-buffered media (Porta et al. 2003, Hassler et al. 2006, Hassler and Twiss, 2006). Further standardization of assay conditions has allowed the use of this strain in documenting Fe bioavailability in the Laurentian Great Lakes (Porta et al. 2003, McKay et al. 2005a, Porta et al.

2005, Hassler et al. 2009). While the use of Fe-responsive bioreporters has offered some evidence of transient Fe deficiency in the Great Lakes (e.g. McKay et al. 2005), it is not widespread (Porta et al. 2005, Hassler et al. 2009). A new approach whereby the bioreporter is contained in a porous underwater chamber (PUC) during incubation with unfiltered water

(Hassler et al. 2008) promises to offer insight into the biological cycling of Fe by regenerative processes such as grazing and viral lysis which may provide 30% to 80% of the algal Fe demand in surface seawater (McKay et al. 2005b).

Kunert et al. (2000) developed luxAB and gfp constructs regulated by the promoter for isiAB in

Synechocystis sp. PCC 6803. As expected, bioreporter luminescence and fluorescence increased in response to Fe deficiency. Both bioreporters also reacted to high salt stress, a response likely related to oxidative stress which appears to be a common control on isiAB transcriptional regulation in cyanobacteria (Michel and Pistorius 2004). 14

The repertoire of cyanobacterial Fe-responsive bioreporters was recently expanded by development of a luminescent whole-cell cyanobacterial bioreporter, strain BMB04 (deposited as

Synechococcus sp. CCMP 2669), constructed using the euryhaline Synechococcus sp. PCC 7002 for measuring Fe availability in diverse marine environments (Boyanapalli et al. 2007). A dose- response curve was generated relating bioreporter luminescence to the free ferric ion content of defined growth medium between pFe 19.4 to 22.4 (see Fig. 1 of Boyanapalli et al. 2007).

Through this range of variable [Fe3+], discernible changes were measured in the luminescent response of cells with luminescence being > 2
higher associated with cells growing under low

Fe (pFe 22.4) compared to Fe sufficient conditions (pFe 19.4). Luminescence plotted as a function of pFe could be described according to a 3-parameter sigmoidal curve and as such, provided a similar response to that which we have characterized for a suite of freshwater Fe bioreporters (e.g. Porta et al. 2003).

Bioreporters of N availability

The bioreporter approach can be viewed as an alternative method to measure nitrogen uptake in aquatic systems. With this rationale, a Synechocystis sp. strain PCC 6803 bioluminescent reporter was developed capable of assessing nitrate assimilatory capacity in freshwaters

(Ivanikova et al. 2005). The bioreporter, designated AND100, is based on the nitrate/nitrite- activated nirA promoter (regulating expression of genes encoding nitrite reductase) which is under positive control by two transcription factors, NtcA and NtcB, that together yield elevated transcription when bioavailable nitrate or nitrite is present in the medium (Frias et al. 2000; Aichi et al. 2001). Characterization of this strain demonstrated a dose-dependent increase in 15

bioluminescence coincident with increased nitrate added to the growth medium to 100 μM.

Additionally, bioluminescence in response to nitrate addition was light dependent up to 50 μmol

quanta m–2 s–1. Because nitrate concentrations in most freshwater systems exceed those of

nitrite by more than an order of magnitude (e.g. Mortonson and Brooks 1980), the bioreporter

can be viewed primarily as a sensor for nitrate bioavailability.

The AND100 strain differs fundamentally from two cyanobacterial nitrogen bioreporters

previously described (Mbeunkui et al. 2002, Gillor et al. 2003). These bioreporters are luxAB

fusions employing the Synechocystis sp. PCC 6803 nblA (Mbeunkui et al. 2002) and S. elongatus

sp. PCC 7942 glnA promoters (Gillor et al. 2003), controlling the genes encoding a

phycobilisome degradation regulator and glutamine synthetase, respectively. Whereas the dynamic ranges of these strains were similar to AND100, the luminescent response was induced upon nitrogen deficiency, not during nitrogen use as shown for AND100. Response time of the S. elongatus sp. PglnA and Synechocystis sp. PnblA bioreporters was also considerably slower, with each reporter strain yielding dose-dependent luminescence on the order of 15 to 25 h (Mbeunkui et al. 2002, Gillor et al. 2003) compared to 4 h for AND100. Additionally, the glnA strain yielded dose-dependent responses to a wide variety of N species ranging from nitrate, ammonium, urea, and glutamine (Gillor et al. 2003), and nblA expression was responsive to nitrate and ammonium

(Mbeunkui et al. 2002). Such broader spectrum responses may be useful properties for the measurement of total nitrogen bioavailability, and indeed, the PglnA reporter has been used to document decreasing nitrogen bioavailability along a west-to-east transect in Lake Erie (Wilhelm et al. 2003). 16

By contrast, the nitrate/nitrite specificity of the AND100 bioreporter provides a means for discriminating between specific nitrogen species. In this context, the AND100 bioreporter likely could provide further insights by focusing on the potential for nitrate use in the Great Lakes.

Nitrate has been accumulating in the Laurentian Great Lakes for decades. Lake Superior represents an end member in this regard with a century-long 6-fold increase resulting in present- day nitrate accumulation approaching 30
mol L-1 (Sterner et al. 2007). Whereas seasonal drawdown of nitrate is demonstrated, it is modest (2-4
mol L-1) compared to the lower lakes in the Great Lakes system (Sterner et al. 2007). To gain insight into the factors that constrain nitrate consumption in Lake Superior, the positive induction of the AND100 luminescent response was used to measure the onset of nitrate use as light levels were manipulated and nutrients (P, Fe) were amended to samples collected from the lake (Ivanikova et al. 2007). These data suggest that

P- and Fe-limited cyanobacteria are deficient in their ability to assimilate nitrate in Lake Superior

(Fig. 3A). Furthermore, during spring, light fluxes are sufficiently low to prevent maximal nitrate utilization, even in the absence of nutrient limitation (Fig. 3B). By comparison, the properties of the Synechococcus sp. PglnA bioreporter, whose bioluminescence is under repression by elevated nitrogen, would not be suitable for such an experiment. Overall, the AND100 strain affords a direct method for determining the role of both chemical and physical factors in regulating nitrate uptake by photoautotrophs.

Whereas the aforementioned strains were developed expressly as environmental sensors, manipulation of cyanobacteria to produce luminescent transcriptional reporters to elucidate components of regulatory networks is a strategy that has been adopted previously (reviewed by

Koksharova and Wolk 2002). Of relevance to N assimilation, Wolk et al. (1991) used a derivative of transposon Tn5 with the filamentous, diazotroph Anabaena sp. PCC 7120 to 17

generate transcriptional fusions to promoterless bacterial luciferase genes. Using this approach, they identified genes that responded to removal of fixed nitrogen by monitoring the

luminescence of colonies from transposon-generated libraries. Likewise, luminescent

transcriptional reporters have been developed to study NtcA- (Aldehni et al. 2003) and NtcB-

regulated expression (Maeda et al. 1998) of genes involved in N assimilation in S. elongatus sp.

PCC 7942. Application of these strains as environmental sensors is a reasonable extension.

Bioreporters of P availability

Phosphorus has been described as the “staff of life – the most essential of nutrients” owing to its importance to biota and the low ambient concentrations at which it is present in many surface waters (Karl 2000). Phosphorus is an essential component of DNA, ATP and phospholipids and

accounts for about 2–4% of the dry weight of most cells (Karl 2000). Yet in many aquatic

environments, inorganic phosphate (DIP) is near detection limits using standard analytical

methods.

Issues and challenges in assessing bioavailable P

Whereas the limitation of primary production by P availability is a central tenet of modern day

limnology, it has only recently been afforded more widespread attention among oceanographers

(see review by Benitez-Nelson 2000). This stems, in part, from a long-held belief that P limits

production only over prolonged time scales. Over shorter periods, N is thought to constrain

phytoplankton growth in the open ocean. This is supported by constraints imposed on 18

phytoplankton nutrient requirements by the canonical Redfield ratio and consideration of ratios

of inorganic N (DIN): DIP that show exhaustion of N before P (Karl et al. 1999). Neglected in

such a consideration, however, is the potential availability of organic nutrients to phytoplankton.

Indeed, when total dissolved P (TDP) is plotted against total dissolved N (TDN) and the trend is

extrapolated to nutrient exhaustion, TDP is depleted before TDN (Karl et al. 1999). This is

important when considering that dissolved organic phosphate (DOP) represents the dominant

fraction of TDP in the upper oligotrophic ocean (Karl and Yanagi 1997, Wu et al. 2000).

Contributing further to our lack of understanding of P bioavailability are the somewhat arbitrary

methods used to measure P (see review by Benitez-Nelson 2000). Most commonly used is the

acid molybdate method capable of measuring P to 30 nM. A modification of this approach, the

magnesium-induced coprecipitation (MAGIC) procedure, is the most sensitive chemical

technique, detecting P to < 5 nM (Karl and Tien 1992). Although long assumed that the acid

molybdate method measures orthophosphate, it is now recognized that formation of the phosphomolybdate complex during sample acidification is accompanied by hydrolysis of some fraction of the DOP pool. Thus, it is more appropriate to refer to the fraction measured using this technique as soluble reactive phosphorus (SRP). To measure TDP, samples are exposed to high temperature and/or pressure in the presence of a strong oxidizing agent prior to phosphomolybdate complex formation. DOP can then be inferred as the difference between

TDP and SRP. Still, interpretation is not straightforward since the presumed DOP fraction may contain some non-reactive inorganic species such as polyphosphates. Alternatively, some components of the DOP fraction may be refractory and are not readily converted to TDP. For example, naturally-occurring phosphonates comprise up to 25% of the high molecular weight 19

DOP pool in the open ocean (Clark et al. 1998), and are generally thought to represent a P source

more refractory to assimilation than organic monophosphate esters (Benitez-Nelson 2000).

Overall, it is clear that current techniques to measure P provide little information on speciation.

Besides the conventional colorimetric methods, several alternative strategies of P determination

based on ion-selective electrodes or enzyme sensors (see review by Engblom 1998) have been

developed in recent decades. Complementary to these approaches are bioassays to assess the

utilization of known or unknown dissolved P species (see review by Boström et al. 1988). One of

the widely used methodologies is the (Provisional) Algal Assay Procedure (as described in

Boström et al. 1988) whereby a P deficient monoalgal culture is inoculated in a sample and algal

yield recorded after several weeks. The yield is converted to P equivalents by calibration with

parallel trials run with DIP although this approach suffers from many of the complications

associated with experiments requiring prolonged incubations.

Many bacteria and phototrophic plankton can exploit forms of DOP, mainly through enzymatic

reactions at the cell surface. Fluorometric ectoenzyme assays offer modest resolution between

DOP species and afford a quantitative method to measure P availability. Two major enzyme

classes appear to be most important in this capacity: alkaline phosphatase, an inducible

monophosphate esterase having broad substrate specificity (Hoppe, 2003) and 5’-nucleotidase, an enzyme capable of hydrolyzing the carbon moiety of nucleotides, releasing DIP in the process

(Ammerman and Azam 1985). A modification of the bulk alkaline phosphatase approach known as enzyme-labeled fluorescence (ELF) affords cell-specific detection of P stress in mixed populations (Dyhrman et al. 2002, Lomas et al. 2004). 20

Additional biochemical and molecular approaches that have been employed include monitoring

the expression of the high affinity periplasmic P-transporter PstS in P-deficient marine

cyanobacteria both by immunoblotting (Scanlan et al. 1997, Scanlan and Wilson 1999, Fuller et

al. 2005) and by reverse transcriptase (RT)-PCR using pstS-specific primers (Dyhrman and

Haley 2006). Recently, the expression of genes involved in phosphonate transport has been

demonstrated in laboratory and field populations of the marine diazotroph Trichodesmium

(Dyhrman et al. 2006) and among picocyanobacteria (Ilikchyan et al. 2009) thus countering the

prevailing view that phosphonates are refractory sources of P.

P-Responsive bioreporters

The use of compound-specific P bioreporters may offer a unique approach toward not only

assessing the bioavailability of various P chemical species but also afford the opportunity to

readily characterize the components of the P pool, especially DOP species at a given site. At

present, investigators are constrained in their ability to characterize the DOP pool by the requirement to concentrate large volumes of water using techniques such as tangential flow ultrafiltration or lyophilization followed by 31P nuclear magnetic resonance (e.g. Clark et al.

1998). By exploiting specific genetic mechanisms used by cyanobacteria to target individual components of DOP, bioreporters offer a means by which to characterize the DOP pool using small volumes of water (as little as 100-200 μL if adapted to a microtiter plate assay). Moreover,

since bioreporter response can be calibrated using model DOP compounds, the concentration of

specific DOP classes in natural waters can be determined. 21

Applicable to monitoring PO4-Pi deficiency in aquatic environments, a luminescent

cyanobacterial P bioreporter was designed by Gillor et al. (2002). This bioreporter, appropriate

for use in freshwater environments, was constructed using a PphoA::luxAB construct integrated

into the cyanobacterium Synechococcus elongatus PCC 7942 and features a detection range of

0.3 – 8.0 μmol L-1 PO4-Pi using a sample incubation time of 8 h. The bioreporter was designed using the promoter for phoA, a P-responsive gene that encodes alkaline phosphatase. The

resulting strain responds by dose-dependent light emission to a wide range of PO4-Pi

concentrations. This freshwater P bioreporter has also been adapted for use as an immobilized

sensor in microtiter plates called the “Cyanosensor” (Schreiter et al. 2001).

Effort is currently being allocated to developing compound-specific P bioreporters in both

Synechococcus sp. PCC 7002 (Nazarov, 2009) and WH8102 (this thesis) able to resolve DIP and

DOP, including nucleotides and phosphonates as P sources. The suite of P bioreporters developed should have the ability to discriminate between use of DIP and DOP and should afford resolution of major classes of DOP utilized by aquatic photoautotrophs.

This thesis describes the construction and initial characterization of two P responsive bioreporter strains for marine environments. These bioreporters were constructed using the promoter regions of two P-stress responsive genes, phoD and nucA , resulting in the strains OPD and OPN respectively.

22

Methods and Materials

Media and growth conditions

Synechococcus sp. WH 8102 was cultured in SN medium (Waterbury et al, 1986) using synthetic ocean water (Price et al., 1988) filter sterilized through 0.2 μm nylon membranes at

25°C under continuous lighting of 25 μmol quanta m-2 s-1 without shaking. To genetically- modified cultures of Synechococcus, spectinomycin (100 μg ml-1) was added to maintain plasmid integrity in liquid culture. Pour plating was done as previously described (Brahamsha, 1996) by serially diluting conjugants and adding them to 50 ml of SN medium containing 0.3% (w/v) purified Bacto Agar (Waterbury et al, 1986 ) pre-cooled to 37°C, and containing 100 μg ml-1 spectinomycin. Plates were aged for 1 week at room temperature prior to use to allow for rapid absorption of culture material once applied.

Glass- and plasticware (polycarbonate tubes) used for cell culture were soaked in 10%

HCl for at least 24 h followed by rinsing 4
in purified water (Millipore Milli-Q Grade). Acid washed material was dried in a HEPA filtered laminar flow hood.

To monitor growth of Synechococcus, in vivo chlorophyll (chl) fluorescence was measured using a TD-700 fluorometer (Turner Designs)

Promoters

Genomic DNA was isolated from Synechococcus sp. WH 8102 using a QIAGEN Blood and Tissue kit and following the manufacturer’s instructions. Promoters were amplified from genomic DNA by PCR (Table 1) and then cloned using the TOPO® TA Cloning System (pCR4

TOPO® or pEXP5 CT TOPO®, Invitrogen). Following transformation into Escherichia coli 23

strain DH5
, plasmids were isolated using a QIAprep miniprep kit (Qiagen). Promoter sequences

cloned into TOPO® vector were verified by automated sequencing at the University of Chicago.

Gel extraction of DNA was performed using a QIAquick Gel Extraction Kit (Qiagen).

All enzymes were from New England Biolabs.

Plasmid Construction

The bioreporter construction was done in a two step process with the first step involving

cloning the promoter sequence of interest into a workhorse plasmid, pALPHA. As a template

plasmid for the construction of pALPHA (Figure 1a), the plasmid pMBR was chosen. pMBR is

a promoter probe constructed for Synechococcus sp. PCC 7002 (Boyanapalli 2006; Boyanapalli et al. 2007) which contains a Vibrio harveyi luxAB cassette as well as a spectinomycin resistance gene. pALPHA was constructed by inserting a synthetically designed 44 bp fragment labeled

SNaP3, into the single SacI site in pMBR. The SNaP3 fragment is a pair of complementary oligonucleotides (Invitrogen) (Table 1) containing a NheI restriction site flanked by two SacI

restriction sites. The SNaP3 oligonucleotides were annealed by combining both forward and

reverse primers into a 200 nM solution, then mixing 1 μg of each primer with 2 μl T4 DNA

ligase buffer and 13 μl sterile water. The reaction mix was incubated at 93°C for 3 min, allowed

to cool to room temperature and used immediately. SNaP3 and pMBR were digested separately

with SacI for 60 min at 37°C and pMBR was incubated for an additional 15 min after adding

Antarctic Phosphatase(New England Biolabs) to prevent ligation of the pMBR without an insert.

Both tubes were then incubated at 65°C for 20 min to deactivate the enzymes. The linearized plasmid and the SNaP3 fragment were ligated for 10 min and transformed into commercially 24

available competent E. coli cells (Invitrogen). Transformed cells were spread on pre-warmed

(37°C) LB plates containing 100 μg ml-1 spectinomycin.

Promoters were fused upstream of the luxAB genes in pALPHA at the BamHI and NotI

sites (Fig 1a). A simultaneous digestion with NotI-HF and BamHI-HF was performed on the

TOPO vectors containing the promoters and on pALPHA for 15 min at 37°C following which

pALPHA was treated with Antarctic Phosphatase. Digestion was verified by mobility on an

agarose gel and the appropriate bands were extracted and ligated (see above).

Constructs to be maintained extra-chromosomally were then developed by inserting the

specr and luxAB cassette from pALPHA containing either the phoD or nucA promoter into

pRL153 (Figure 1b), an IncQ type incompatibility plasmid derived from RSF1010 (Elhai and

Wolk, 1988). This resulted in the plasmids pOPD (Figure 1c) and pOPN (not shown)

respectively. The extrachromosomal plasmids were constructed by digesting pALPHA and

pRL153 with NheI-HF following which pRL153 was treated with Antarctic Phosphatase.

Digestion was verified by mobility on an agarose gel and the appropriate bands were extracted

from agarose gels and ligated (see above).

Conjugation into Synechococcus

The extrachromosomal plasmids pOPD or pOPN, were transferred into Synechococcus

sp. WH 8102 via conjugation (Brahamsha, 1996). E. coli containing an extrachromosomal construct and the conjugal plasmid pRK24 were grown overnight in LB broth containing spectinomycin, ampicillin, and tetracycline, washed 3
and resuspended in SN medium containing 10% LB. Twenty-five ml of mid-log phase Synechococcus sp. WH 8102 were 25

Table 1. Primers used in this study

Primer Sequence phoD-F Atttgcggccgctttatcagctggccctgatcgatttg phoD-R Cgcggatccgcggatttttcttcagactttgttaattctgg nucA-F Cgcggatccgcgtgtgccgaaagtttcttcataa nucA-R Atttgcggccgctttatcgattttaggttgcacttcc SNaP3-F tgattacggcttcccatctgcgaacgagctcgaaaaaggggaaaaaagctagcggaagggaaaggaaacgagctcgaagtcggtaacggtgtggaagtaa SNaP3-R ttacttccacaccgttaccgacttcgagctcgtttcctttcccttccgctagcttttttcccctttttcgagctcgttcgcagatgggaagccgtaatca prochk-F gccctacacaaattgggaga prochk-R cggaatcgtccttttgacat

Table 2. Plasmids used in this study.

Plasmid Characteristics Reference pRK24 Apr; Tcr; conjugal plasmid Meyer et al, 1977 pRL153 RS1010 derivative; Kanr; contains mob genes Elhai and Wolk, 1988 pMBR pAM1414 derivative; contains luxAB; Specr; Boyanapalli et al, 2007 pALPHA luxAB; Specr; This study pOPD luxAB; Specr; Kanr;phoD promoter This study pOPN luxAB; Specr; Kanr;nucA promoter This study

26

Figure 1. Map of pALPHA, pRL153, and pOP(D) A. pALPHA plasmid is a workhorse plasmid into which promoters of interest are inserted into the NotI and BamHI sites. Contains the luxAB and spectinomycin resistance cassettes. Constructed by inserting the SNaP3 fragment (See methods and

Materials) into a SacI site enabling the luxAB and spectinomycin cassettes to be inserted into pRL153. B. pRL153, an RS1010 derivative IncQ incompatibility plasmid. Contains the mob genes necessary for conjugation, origin of transfer (oriT), the rep genes involved in replication and copy number control, and kanamycin resistance. C. pOPD. This is an extra chromosomal construct created by fusing the luxAB/Spec cassette from pALPHA into pRL153, containing the phoD promoter to be maintained in

Synechococcus spp WH 8102. 27

centrifuged at 5,856
g at 20°C for 10 min and the pellet was resuspended in 150 ml SN

medium. Mixtures were then made in several dilutions by adding 50-, 25- and 10 μl of

resuspended Synechococcus to 50 μl of transformed E. coli. One drop of each dilution was pipetted onto an aged (1 week) SN plate in a single spot and allowed to absorb into the plate.

The plates were maintained at low light (10 μmol quanta m-2 s-1) for 48 h at 25°C. The spots

were cut out using a sterile spatula and resuspended in 2 ml SN medium. These were then

serially diluted and plated as above. Single colonies were pipetted out of the gel and suspended

in 2 ml SN medium. Residual E. coli was tested for by streaking on LB plates and incubating

overnight at 37°C.

Colony PCR was used to verify the insertion of the extrachromosomal construct. The

PCR reaction was set up according to manufacturer’s instructions. For a template, 1 ml of

culture was spun down at 14000 rpm for 10 min, after which the supernatant was discarded. A

sterile pipette tip was touched to the pellet and then touched to the PCR reaction. The PCR

cycle used was as follows: Cycle 1 (
20): 1 min at 94°C, 1 min at 65°C, 1 min at 72°C; Cycle

2 (
20): 1 min at 94°C, 1 min at 55°C, 1 min at 72°C.

Bioreporter Characterization

The P-dependent steady-state luminescent response of both the nucA and phoD

bioreporters was assessed by incubating the bioreporter in SN medium containing defined

amounts of potassium phosphate. Phosphate-depleted cells were grown in SN medium

containing 200 nM P whereas P sufficient cultures were grown in medium containing 80 μM P.

At daily intervals, 2 ml of sample was removed from each culture and growth was assessed by 28

measuring chl fluorescence. P-responsive bioreporter luminescence was measured immediately

following addition of 60 μM n-decyl aldehyde using a portable Femtomaster FB14 luminometer

(Zylux Corp.) with settings of 5 s delay and 10 s measurement. To account for differences in cell

biomass between treatments, luminescence was normalized to chl fluorescence.

Results

Verification of plasmid insertion

Insertion of the extrachromosomal plasmid was verified by performing colony PCR on samples

from the cultures using primers flanking the promoter insert site (prochk-F and prochk-r, Table

1). As controls, colony PCR was performed on E. coli used for conjugation of pOPN and a

plasmid miniprep of the pOPD culture was used for template DNA (QIAprep miniprep kit,

Qiagen). A negative control contained no template DNA in order to test for contaminants.

Analysis on a 1% agarose gel (Fig. 2) showed bands of appropriate sizes of 957 bp and 879 bp

for pOPN and pOPD, respectievly. The negative control showed no amplification of DNA,

consistent with no contamination in the PCR setup. Tests for residual E. coli were negative, ruling out a band generated from the donor strains.

Growth of wild type vs. bioreporter strains

Growth rates of bioreporter strains containing the extrachromosomal construct were higher than for wild type Synechococcus sp. WH 8102 when cultured in P-sufficient SN medium (P<0.001,

one-way analysis of variance followed by Tukey honestly significance difference [HSD]

test)(Fig. 3) . Wild type cells possessed a growth rate of 0.305 ± 0.06 d-1 compared to 0.495 ± 29

0.04 d-1 and 0.538 ± 0.05 d-1 for the bioreporter strains incorporating nucA and phoD, respectively. There was no significant difference (P = 0.464; df = 11) in growth rate between the two bioreporter strains. Subsequent experiments were conducted only with the genetically modified strains.

Delivery of aldehyde substrate to the bioreporter

The bioreporter construct was designed to require exogenous application of decanal substrate for bacterial luciferase since endogenous substrate levels are rarely adequate as produced by strains co-transformed with aldehyde-producing luxCDE genes (e.g. Porta et al. 2003). Whereas some studies, including the prior characterization of a freshwater Fe bioreporter (Porta et al. 2003), have advocated delivery of aldehyde via the vapor phase, commonly, exogenous aldehyde substrate is injected into the medium to elicit a luminescent response (Ivanikova et al. 2005).

Previous efforts from our lab group to use vapors to deliver decanal to the bioreporter cells have consistently provided high variability among replicate samples (R.M.L. McKay, personal communication). As a result, substrate delivery by direct injection was chosen for subsequent characterization of the bioreporter. As part of the present characterization of the Synechococcus sp. WH8102 P-responsive bioreporters, the concentration of n-decyl aldehyde substrate was optimized, and the efficacy of methanol vs. water compared as solvents for delivery of the aldehyde substrate.

Comparing solvents for substrate delivery, aldehyde delivered in pure methanol resulted in lower luminescence readings whereas no difference was recorded when aldehyde was 30

Figure 2. Colony PCR verification of plasmid insertion in Synechococcus spp. WH8102.

PCR was performed on pOPN and pOPD cultures, pOPN E-coli stock, and a plasmid prep of pOPD using the prochk primers. The molecular weights obtained match the expected values of

957 bp (pOPN) and 879 bp (pOPD).

31

Figure 3. Growth of bioreporter strains OPD (closed circle) and OPN (open circle), compared with the growth of wild type Synechococcus spp. WH8102 (closed triangle). The rate

of growth was slower, and the onset of exponential growth delayed, for the WT strain compared

to the two bioreporter strains.

32

delivered using 50% methanol or water alone (P<0.001, one-way analysis of variance followed

by Tukey HSD test) (Fig. 4a). A study by Hassler et al (2006) demonstrated that the addition of

60
mol L-1 n-decyl aldehyde in water gave the highest luminescence which was consistent with our trials. This mode of delivery was adopted for the remainder of this study.

Cell luminescence followed similar kinetics regardless of the n-decyl aldehyde substrate concentration used (Fig. 4b). In each case, luminescence was high immediately following substrate addition, declining thereafter and reaching its lowest value between 1-2 min.

Luminescence then again began to increase with values increasing through the 5 min assessment shown here. Readings beyond this point showed maximum luminescence at around 20 min but only slightly higher than the reading taken immediately following aldehyde addition (data not shown). Regardless of the concentration used, delivery of n-decyl aldehyde resulted in nearly identical cell luminescence kinetics with luminescence recovering to near initial values within 5 min with no significant differences (P<0.001; df = 11, one-way analysis of variance).

P-Responsive Luminescence of Bioreporter Strains

The luminescence of the nucA and phoD bioreporters was characterized in response to steady-state growth in P depleted (200 nM) and P sufficient (80μM) media (Fig. 5). When grown in P sufficient medium, both bioreporters maintained lower luminescence then when grown in P depleted medium. Initially, growth in either P-sufficient or P-depleted medium elicited a similar elevated luminescent response from the bioreporters (Figs. 5 a,b). Differences in P-responsive luminescence for the phoD bioreporter, POPD, were evident following 2 days of 33

Figure 4. Comparison of solvent type and concentration of n-decyl aldehyde concentrations on cell luminescence. A. The effect of solvent type for delivery of aldehyde on cell luminescence.

B. Comparison of n-decyl aldehyde concentrations on cell luminescence. 34

incubation with sustained higher luminescence in P-depleted medium compared to P sufficient

cultures (P < 0.05, t-test) (Fig. 5a). The nucA (POPN) bioreporter followed a similar response;

however, the difference was not evident until after 5 days incubation ( P < 0.05, t-test) (Fig. 5b).

Whereas the growth of bioreporter strains was robust in P sufficient medium, growth was

inhibited in P-depleted medium until P was added to the medium on day 9 (Figs. c,d). Once

growth entered exponential phase in these cultures, cellular luminescence dropped to a level

similar to that associated with P replete cultures, suggesting that both bioreporters were

responsive to P levels in the medium.

Discussion

The cyanobacterium Synechococcus sp. WH8102 presents itself as a viable candidate to host a

nutrient-responsive bioreporter for a variety of reasons: it is a pelagic strain, ecologically

relevant to the open ocean, its genome has been fully sequenced (Palenik et al, 2003), and it has been shown to be amenable to genetic manipulation by conjugation (Brahamsha, 1996).

Moreover, the phosphorous assimilation pathways of Synechococcus sp. WH8102 have been

analyzed in silico (Su et al. 2003).

Promoters on which to base bioreporter construction were chosen from a number of

possible P-responsive genes likely to be indicative of early P stress. Promoters chosen for the

constructs were phoD from the P regulon (Su et al. 2003), encoding a phosphodiesterase - alkaline phosphatase and nucA (Ammerman and Azam, 1985), which encodes a 5
nucleotidase.

A recent study using microarray analysis on P depleted cultures of Synechococcus spp. WH8102 35

Figure 5. Luminescence expression plotted with growth curves for nucA and phoD bioreporters in both P sufficient and -depleted conditions. Arrows indicate the addition of 80 μM potassium phosphate to the cultures. A/B. Luminescence readings normalized to in vivo chlorophyll for the phoD (A) and nucA (B) bioreporters. C/D. Growth curves (ln of in vivo chlorophyll) for phoD (C) and nucA (D) bioreporters.

36

reported strong up-regulation of both phoD and nucA genes during early P stress (Tetu et al,

2009), making them ideal candidates for early P stress detection in a whole cell bioreporter and validating our choice of promoters. The amplified promoter sequence for nucA is the 245 base

pair region located upstream of the SYNW2390 start codon whereas phoD is the 167 base pair

region located upstream of the SYNW0196 start codon in the Synechococcus sp. WH 8102

genome (NCBI).

Alkaline phosphatase, a cell surface enzyme, hydrolyses orthophosphate from dissolved

organic phosphorus (DOP) when dissolved inorganic phosphorus (DIP) is depleted (Perry,

1972). It has been shown that under P deficiency, marine algae synthesize alkaline phosphatase

to levels 30 times or higher than levels measured in P sufficient cultures, and also that its

synthesis is repressed under P-replete conditions (Kuenzler and Perras, 1965). In marine

phytoplankton, alkaline phosphatase is often used as an indicator of P stress. In the Sargasso

Sea, ELF-labeling (enzyme-labeled fluorescence) for alkaline phosphatase was employed to

assess P stress in phytoplankton (Lomas et al, 2004). In that study, marine Synechococcus were

not labeled, suggesting that the cells were not P-deficient.

The enzyme 5
nucleotidase offers an alternative approach to acquisition of P. As cells

lyse, they release nucleic acids into the environment. It has been suggested that exploitation of P

contained in nucleic acids using 5
nucleotidase, could supply up to half of the P required by

cyanobacteria (Ammerman & Azam, 1985). Consistent with this, 5
nucleotidase activity has

been reported associated with marine cyanobacteria and measured in diverse marine and

estuarine environments (Ammerman & Azam, 1991).

The workhorse plasmid, pALPHA (Figure 1a), was designed for the construction of the

extrachromosomal plasmid pOP used in this study as well as for more facile construction of 37

future bioreporters. The SNaP3 fragment was inserted downstream of the luxAB cassette to

complement the NheI site upstream of the spectinomycin cassette, which allows for easy removal

of the entire fragment containing the spectinomycin and luxAB cassettes, along with the promoter of interest, by digestion with NheI. The entire fragment can then be inserted into any plasmid containing a single NheI site. The use of a workhorse plasmid is useful for this application since the same promoter and cassette can be inserted into a new plasmid construct, such as a future integrative plasmid, with minimal effort. The plasmid chosen as the host plasmid for the extrachromosomally-maintained bioreporters was pRL153 (Figure1b). This plasmid is an IncQ

incompatibility plasmid containing a kanamycin resistance cassette as well as the mob genes

necessary, in conjunction with pRK24, to mobilize the plasmid into E. coli. The resulting

bioreporter, OPD or OPN in this study, thus contains a luxAB cassette containing a promoter of

interest as well as both spectinomycin and kanamycin resistance. In addition, it has mobilization

properties for conjugation into Synechococcus WH 8102.

The prochk primers (Table 1) were designed to anneal to regions upstream and

downstream of the promoter region, amplifying a region of 698 bp plus the length of the inserted

promoter. These primers can be used to test the insertion of any plasmid containing this

lux/spec cassette from pALPHA.

The initial characterization of both OPD and OPN bioreporters show that they are

responsive to P limitation in synthetic media, with luminescence increasing in P-depleted

cultures and decreasing with the addition of 80 μM potassium phosphate. OPN and OPD appear

to have similar response time after the addition of P, both reaching luminescence levels of the P-

sufficient culture within 6 days. When comparing the initial response to P depletion, OPD 38

responded faster than OPN, with a separation from P sufficient samples within 3 days whereas

OPD did not differ from P sufficient samples until day 6. This is at odds with microarray data which suggests that nucA responds faster to P stress than does phoD (Tetu et al, 2009). The delay in both strains may be due to damage to the cells during centrifugation or possibly a result of P stress before inoculation in fresh medium. In comparison with the freshwater P bioreporter strain APL, the OPD and OPN strains first appear to have a slower reaction time, taking days to differentiate between P stressed and P sufficient samples, whereas APL can differentiate between

P stressed and P sufficient samples within hours (reference needed). On the other hand, after differentiating from P sufficient samples, OPD reaches maximum recorded luminescence within

2 days and OPN reaches maximum luminescence within 24 h, whereas APL can reach maximum luminescence in less than two days, suggesting that once the right starting conditions are attained, OPN and OPD may have similar kinetics to APL.

Conclusion

Picocyanobacteria are dominant primary producers in many pelagic waters, and therefore an excellent indicator of the integrity of an environment. This highlights their functionality in identifying and monitoring factors constraining productivity. Whereas inorganic P can be measured using a variety of chemical tests, the potential use of organic P is often negelected.

The OPN and OPD bioreporters were designed to complement existing methods of P measurement, as well as to provide an ecologically relevant bioreporter for the open ocean.

These reporters show the feasibility of Synechococcus spp. WH8102 as a bioreporter strain. 39

Initial characterization of this strain demonstrates its potential use in detecting bioavailable P, as well as providing a near-real time view of the environment from a microorganism’s perspective.

40

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