T7 RNA Polymerase Functions In Vitro without Clustering
Abstract 
Many nucleic acid polymerases function in clusters known as factories . 
We investigate whether the RNA polymerase ( RNAP ) of phage T7 also clusters when active . 
Using ` pulldowns ' and fluorescence correlation spectroscopy we find that elongation complexes do not interact in vitro with a Kd ,1 mM . 
Chromosome conformation capture also reveals that genes located 100 kb apart on the E. coli chromosome do not associate more frequently when transcribed by T7 RNAP . 
We conclude that if clustering does occur in vivo , it must be driven by weak interactions , or mediated by a phage-encoded protein . 
Introduction
Mounting evidence suggests that many RNA and DNA polymerases function in clusters rather than in isolation . 
Mammalian RNA polymerase II ( RNAP II ) , for example , appears to be active in ` factories ' which typically contain ,8 enzymes working on different templates , and DNA polymerases cluster in analogous ` replication factories ' [ 1,2,3 ] . 
Such ` factories ' may also exist in 
The single-subunit RNA-dependent RNA polymerases of many human viruses also cluster , forming large membrane-bound arrays in which individual molecules interact directly [ 8,9 ] . 
The formation of these assemblies can have strong effects on RNAP function ; poliovirus RNA-dependant RNAPs , for example , can not transcribe efficiently without forming clusters [ 10 ] . 
Although there are many ways in which the cell might benefit from the existence of polymerase clusters [ 1 ] , the evolutionary forces responsible for their formation remain poorly understood . 
One possibility is that clustering creates a high local concentration that facilitates nucleic acid synthesis [ 11 ] . 
Another is that RNAP clustering evolved because freely-mobile enzymes would track along and rotate about their templates , and so entangle their trailing nascent transcripts ; conversely , RNAPs immobilized in clusters would reel in their templates without rotating , and so extrude unentangled transcripts [ 11 ] . 
The RNAP of bacteriophage T7 is one of the best studied DNA-dependant RNAPs . 
The conformation of this single-subunit enzyme remains largely unchanged during promoter binding and polymerization of the first three nucleotides [ 12,13,14 ] ; however , by +7 , the enzyme has already undergone significant rearrangements [ 15 ] and by +14 has morphed into its final processive form [ 16,17 ] . 
The resulting elongation complex ( EC ) is highly stable [ 18 ] , and transcribes at ,50 -- 200 bp/s [ 19,20 ] . 
Little is known about the clustering of any of these T7 RNAP isoforms . 
However the unengaged enzyme does ` aggregate ' at the high concentrations ( ,10 mM ) used during purification and crystallization [ 21,22,23 ] -- and so is often solubilized using nonphysiological concentrations of NaCl and glycerol [ 24,25 ] . 
It is not known whether this interaction is physiologically relevant , or occurs at lower RNAP concentrations . 
Whether ECs cluster is equally unclear . 
Although isolated monomers can function when immobilized in vitro [ 19,26 ] , it remains to be seen whether ECs cluster in vivo or in solution . 
ECs have been imaged by atomic force microscopy and appear as monomers [ 27 ] ; however , the procedures used to prepare these samples may have destroyed any pre-existing clusters . 
Here , we investigate whether or not T7 RNAP ECs cluster using ` pulldowns ' , fluorescence correlation spectroscopy , and chromosome conformation capture . 
We find no evidence for clustering , and conclude that if it does occur in vivo , it is probably driven by weak interactions . 
Results
T7 RNAP ECs do not co-associate in vitro
To test whether active T7 RNAPs cluster , we examined whether ECs diffusing freely in solution interacted with distinguishable ECs directly attached to beads ( Fig. 1A ) . 
To achieve this , we created a transcription reaction containing RNAP as well as three DNA fragments of different lengths ( Fig . 
S1A ) : a 290-bp template encoding a T7 promoter that was freely-diffusing in solution , a 452-bp template which again encoded the promoter but was bound by a biotin at its 59 end to streptavidin-coated present in samples 1 -- 3 , but not 4 ( as it fails to pellet ) . 
The 452-bp template is present in samples 1 and 4 ( as it binds to beads , and pellets ) . 
Only trace amounts of the 290-bp template migrate as free DNA in sample 2 ( elongation complexes migrate more slowly as a smear ) , but this amount is increased in sample 3 ( as RNase and heat treatments release it from elongation complexes ) . 
The 290-bp template is found in sample 4 when the assay is performed in 10 mM KCl . 
However it is absent when the assay is performed in 10 mM KCl plus tRNA , or the more physiological buffer containing 100 mM K glutamate . 
doi :10.1371 / journal.pone .0040207 . 
g001 beads , and an 800-bp promoter-less control fragment . 
When ATP , UTP , and GTP ( but no CTP ) were added , RNAPs initiated on the two templates encoding promoters , and transcribed until they needed to incorporate CTP ; they then stably halted ( Fig . 
S1B ; previous work has shown that the resulting halted ECs have half-lives .10 min ; [ 18 ] ) . 
We then isolated the ECs formed on the 452-bp templates by pelleting the beads and removing the supernatant . 
Any ECs formed on 290-bp templates interacting with these pulled-down ECs would then be found in the pellet . 
When the pelleted DNA was isolated and visualized , a small amount of the 290-bp template -- but virtually no 800-bp control DNA -- was found ( Fig. 1Bi , sample 4 ) . 
Thus it seemed that ECs on the 290-bp template were associating with the beads and being pelleted . 
Examination of the DNA remaining in the supernatant using agarose gel electrophoresis allowed us to distinguish unbound templates ( which migrate as free DNA ) from occupied templates ( which migrate more slowly ; Fig . 
S2 ) . 
When the RNAPs in the removed supernatant are stripped from their templates ( by heating ) before gel electrophoresis , a large amount of 290-bp template migrates as free DNA ( Fig. 1Bi , sample 3 ) . 
However very little 290-bp template migrates freely when RNAPs remain bound to their templates ( Fig. 1Bi , sample 2 ) . 
These results suggest that the majority ( i.e. , 60 -- 80 % ) of 290-bp templates were occupied by halted RNAPs at the moment the beads were pelleted . 
Additional controls showed that RNAPs initiated as efficiently on the 452-bp template as on the 290-bp template ( Fig . 
S3 ) . 
Thus , we conclude that although the majority of 452-bp and 290-bp templates were occupied by RNAPs , only a small fraction of the 290-bp was pelleted . 
However , we were concerned that the interaction between ECs might be caused by aggregation of nascent RNA , and not by an interaction between RNAPs . 
To investigate this possibility , we repeated the experiment in a buffer containing 10-fold more tRNA than DNA template ( Fig. 1Bii ) . 
We expected that the tRNA would disrupt any non-specific RNA-based interactions ( by competing for any RNA-binding sites ) , while leaving polymer-ase-based protein-protein interactions unaffected . 
When the experiment was conducted in the presence of tRNA , only tiny amounts of the 290-bp template were found in the pellet ( `` 8 % of total ; Fig. 1Bii , compare samples 4 and 5 ) . 
Because the remaining 290-bp template did not appear to be enriched relative to the 800-bp promoter-less control fragment ( Fig. 1Bii , compare samples 4 and 5 ) , we concluded it was not pelleted due to EC-EC interactions , but rather , persisted because we only removed ,97 % of the supernatant . 
Our finding that no short template ( or control DNA ) was found in the pellet when a gentle wash step was included supports this interpretation ( data not shown ) . 
Therefore , we conclude that the previously-observed interaction was based on non-specific RNA interactions . 
As such interactions are unlikely to be physiologically relevant ( see Text S1A ) , we conclude that no meaningful RNAP-RNAP interactions were detected using these assay conditions . 
Repeating the assay using a more physiological buffer ( KGB , which contains 100 mM K glutamate , instead of LS1 , which contains 10 mM KCl ) yielded a similar conclusion even though no tRNA was present : although most templates were occupied by RNAPs ( Fig. 1Biii , compare free-migrating short template in samples 2 and 3 ) , no enrichment of the 290-bp template relative to the control DNA was observed ( Fig. 1Biii , sample 4 ) . 
Identical results were obtained when the total concentration of ECs was increased to 0.1 mM , and when bovine serum albumin was used as a blocking agent instead of casein ( data not shown ) . 
Were ECs to form stable , oligomeric clusters , we would expect that most of the occupied short template ( i.e. , ,60 -- 80 % of total ) would interact with the bead-bound ECs , and so be found in the pellet . 
Our finding that less than a few percent of the short templates are pulled down therefore supports the conclusion that ECs do not form stable clusters under these conditions . 
T7 RNAP ECs do not interact with a Kd ,1 mM In our previous experiment , we found that ECs attached to beads were unable to ` pull down ' ECs in solution . 
However , it is possible that the pelleting of the bead-bound ECs disrupted their interaction with ECs in solution . 
To eliminate this possibility , we used fluorescence correlation spectroscopy ( FCS ) to study EC diffusion behaviour . 
In this nonperturbative technique , a laser is focused on a ` confocal spot ' in solution , allowing the measurement of the diffusion times -- and therefore relative sizes -- of fluorescently-labelled ECs [ 28 ] . 
Since diffusion is slower for larger complexes , diffusion times increase with complex size . 
We expected single ECs with no interaction partners to diffuse relatively quickly , with a small diffusion time less than or equal to the sum of the diffusion times of their components ( i.e. , an RNAP and its template ; Text S1B ) ; in contrast , interacting ECs should diffuse more slowly as large complexes containing multiple RNAPs and templates -- with diffusion times greater than those expected for non-interacting ECs . 
We began by calculating an expected diffusion time for noninteracting ECs . 
We determined that the diffusion time of the 70-bp fluorescently-labeled template upon which our ECs would be formed was 2.460.1 ms ( Fig. 2Aii ) . 
This measurement was in agreement with values determined previously ( Text S1C ) . 
We then calculated that T7 RNAP would -- because of its size and globular nature -- have a diffusion time of 2 -- 3 ms ( Text S1C ) . 
Assuming that the diffusion time of a complex would be less than the sum of the diffusion times of its parts , we concluded that non-interacting ECs would have a diffusion time of 2.4 -- 5.4 ms. If ECs had a diffusion time above this range , it would suggest the existence of larger , and therefore higher-order , complexes . 
To generate ECs that could be tracked by FCS , we allowed RNAP to initiate on a 70-bp fluorescently-labeled template in the presence of ATP , UTP , and GTP . 
Under these conditions , the enzyme produced a 23-bp transcript before stably halting when the first C needed to be incorporated ( Fig . 
S1 ) . 
The majority of such a short nascent transcript is hidden within the RNAP ( or bound to its surface ; [ 27 ] ) , and we anticipated that the few bps emerging from the EC would not drive the RNA-based interactions observed in our ` pulldown ' assay . 
We expected that the templates in the EC-containing solution would be found in one of three populations : unoccupied templates , templates incorporated into ECs that are not bound to other ECs , and templates incorporated into ECs which in turn are bound to other ECs . 
For complexes with diffusion times within an order of magnitude of one another , FCS essentially reports the average diffusion time of all fluorescent species ; thus fast-diffusing templates not bound to clustered RNAPs could -- if numerous enough -- easily obscure the existence of more slowly-diffusing EC clusters . 
To ensure that the fraction of templates not incorporated into ECs was negligible , we used more RNAP than template in our reactions , and performed extensive controls to show that virtually every template was bound by an active RNAP ( Text S1D ) . 
The fraction of ECs found in clusters depends upon the strength of the attraction between RNAPs ; as most protein-protein interactions have Kd between 1 nM and 1 mM [ 29 ] , we expected that the strength of any EC clustering would also fall within this range . 
To detect such interactions , we required EC concentrations .0.1 mM ; unfortunately , our FCS setup could only measure fluorescent species present at concentrations below 50 nM . 
To allow higher concentrations of ECs , we used a low concentration of labeled template ( always 2 nM ) and a large excess of unlabeled template ( up to 0.54 mM ) in our transcription reactions . 
ECs formed on unlabeled templates would not be directly visible to our FCS assay , but could still bind to the labeled ECs and so retard their diffusion . 
After initiating a transcription reaction containing 2 nM labeled 70-bp template , 100 nM unlabeled 70-bp template , and 120 nM RNAP , we measured the average diffusion time of the now-occupied templates to be 3.360.2 ms ( Fig. 2Aiii ) . 
To be absolutely confident that all templates were incorporated into ECs ( Text S1D ) , we repeated the experiment using an increased RNAP : template ratio of 5:1 ; the template diffusion time marginally increased to 3.960.2 ms ( Fig. 2Aiii ) . 
These diffusion times fall squarely within the range expected for non-interacting ECs , and thus provide no evidence for RNAP clustering . 
However , we were unable to calculate precisely an expected diffusion time for small EC clusters ( e.g. , dimers or trimers ) , and thus could not formally exclude the possibility that our ECs were diffusing as dimers or other lower-order complexes , rather than monomers . 
To set a lower limit on the diffusion times of EC clusters , we replaced the 70-bp unlabeled templates in our experiment with 452-bp unlabeled templates ( Fig. 2Aiv ; S1 ) . 
Under these conditions , any EC clusters would contain at least one EC formed on a 452-bp template , and so would possess a D. 15 ms ( i.e. , the diffusion time of the 452-bp template alone ; Fig. 2Av ; Text S1C ) . 
However , substituting unlabeled 452-bp templates for unlabeled 70-bp templates had no significant effect on the diffusion time of the labeled 70-bp ECs , which still diffused with D = 3 -- 4 ms ( Fig. 2Aiii -- iv ) . 
This was the case even when the concentration of occupied 452-bp templates was increased to 0.54 mM ( Fig. 2Aiv ) . 
We conclude that -- under our assay conditions -- the overwhelming majority of RNAPs halted on the labeled 70-bp templates did not bind to the RNAPs halted on the 452-bp templates . 
We note that our finding that the diffusion times of ECs was relatively unaffected by the ratio of RNAP : template is not consistent with the possibility that an interaction was present , but titrated out by excess RNAP . 
To estimate the detection limit of our assay , we calculated the autocorrelation function that our assay would have produced , if the halted RNAPs were to interact . 
In the experiment of Figure 2Aiv , we measured the autocorrelation function of 2 nM labeled ECs ( formed on 70-bp templates ) , in the presence of 0.54 mM unlabeled ECs ( formed on 452-bp templates ) . 
If ECs dimerized with Kd = 1 mM , such a solution would contain ,40 % dimers and ,60 % monomers . 
We calculated the autocorrelation function of this solution by conservatively modeling monomers ( 70-bp templates bound by halted RNAPs ) as having a tD of 4 ms , and dimers ( complexes containing two active RNAPs , one 70-bp template , and one 452-bp template ) as having a tD of 15 ms. We find that such a solution would produce an autocorrelation function clearly distinguishable from the one measured in the experiment summarized in Fig. 2Aiv ( with results in Fig. 2B ) . 
Thus , we conclude that -- under our in vitro conditions -- active T7 RNAPs do not interact with a Kd ,1 mM . 
Genes transcribed by T7 RNAP do not detectably interact 
To test whether ECs interact in their native cellular environment ( i.e. , in living E. coli ) , we used ` chromosome conformation capture ' ( 3C ; [ 30 ] ) to determine whether or not two T7 promoter-encoding genes -- which are located far apart on the bacterial chromosome -- are in contact more frequently when transcribed by T7 RNAP . 
If ECs active at different genomic sites interacted , we expected that their respective transcription units would also be brought into close proximity . 
We began by constructing a strain that would allow us to test this hypothesis . 
We first inserted two genes encoding T7 promoters ( PT7-YFP and PT7-T7gene10 ) into the E. coli genome 100 kbp apart ( Fig. 3A ) . 
We expected that if ECs clustered , these two genes would be brought into contact when transcribed by the T7 polymerase . 
To control the levels of T7 RNAP in the cell , we integrated a gene expressing the polymerase under the control of a PBAD promoter ( Fig. 3A ) . 
This gene produced high levels of T7 RNAP when cells were grown in arabinose , but negligible levels when cells were grown in glucose ( Fig. 3Bi ) . 
Controls confirmed that this T7 RNAP efficiently transcribed the two T7 promoterdriven test genes ( Fig. 3B ) . 
We then used ` 3C ' to determine whether or not the two testgenes were in contact more frequently when transcribed by T7 RNAP . 
This PCR-based method determines the relative interaction frequencies of different genomic regions in vivo [ 30 ] . 
Cells are fixed with formaldehyde , and their chromatin digested with a restriction enzyme . 
Cross-linked restriction fragments are then ligated together , and the frequency of ligations between different pairs of restriction fragments is measured by PCR . 
We performed 3C on cells grown in either arabinose or glucose , and -- under both conditions -- determined the frequency with which the BglII restriction fragment containing PT7-T7gene10 was ligated to the fragment containing PT7-YFP ( Fig. 4A ) . 
We found that transcription of the two test-genes by T7 RNAP had no effect on the ligation frequency of their respective restriction fragments ( Fig. 4B , lanes 1,2 , primer pair a : c ) . 
Controls showed that the formation of the ligation products depended on formaldehyde crosslinking ( Fig. 4B lane 3 ) , and that the efficiency of the 3C protocol was independent of the presence of T7 RNAP ( Fig. 4B primer pairs a : b , d : e ) . 
We conclude that if T7 RNAP ECs do interact , they do not do so strongly enough to significantly change 
Discussion
Many RNAPs co-associate when active ; this clustering often influences function , for example , by increasing activity ( see Introduction ) . 
In order to determine whether T7 RNAP behaves similarly , we used three independent assays to test whether this polymerase also clusters when active . 
In the first assay , we attempted to ` pulldown ' ECs in solution using ECs attached to beads ( Fig. 1A ) , and found no evidence for a direct protein-protein interaction ( Fig. 1B ) . 
As this assay required physical manipulation of ECs which might break weak EC-EC interactions , we performed a second assay using fluorescence correlation spectros-copy ; this directly measures complex sizes without the need for physical manipulation , but it also failed to provide evidence for clustering ( Fig. 2 ) . 
Therefore , if T7 ECs do interact in vitro , it seems likely that they will do so with a Kd outside the detection range of our assays ( i.e. , .1 mM , which is much greater than the estimated in vivo concentration of 30 nM ; see Text S1G ) . 
As the buffers and enzyme concentrations we use are typical of those widely applied by others [ 18,20,24 ] , we conclude that in the majority of the instances where it has been studied , T7 RNAP has behaved as a monomer . 
Because interactions present in vivo can be missed by in vitro assays ( e.g. , if they require macromolecular crowding , or a ` bridge ' protein ) , we also used chromosome conformation capture ( 3C ) to examine association in vivo ( Fig. 3 ) . 
In mammals , 3C readily detects RNAP-driven clustering of active genes [ 31,32 ] , even when those interactions occur in only ,1 % cells in the population [ 31 ] . 
However , 3C failed to provide any evidence for clustering in bacteria ( Fig. 4 ) , even though the genes we examined are probably as tightly packed with polymerases as the ribosomal cistrons ( our T7 RNAP-based expression system can produce as much RNA as all seven ribosomal cistrons combined , which are each typically occupied by 70 RNAPs/gene ; [ 33,34 ] ; see also Text S1E ) . 
However , our 3C assay does have limitations . 
It involves formaldehyde fixation , which can rapidly disrupt nucleoid structure [ 35,36 ] , and so could -- in principle -- also destroy any clustering . 
Note , however , that clustering of genes binding H-NS , a global transcriptional silencer , can be detected by 3C [ 37 ] . 
We may also have inadvertently inserted our two test genes in regions of the bacterial genome that interact rarely . 
Another problem is that the phage-encoded proteins expressed during T7 infection were not present in our 3C assay . 
Any EC clustering dependent upon a phage-encoded ` bridge ' protein would not have been detected in our assays ( this , and other potential problems are discussed in Text S1F ) . 
In conclusion , we find no evidence for the clustering of active forms of T7 RNAP either in vitro or in vivo . 
Our in vitro assays allow us to exclude the possibility of a strong interaction between ECs ( i.e. , with Kd ,1 mM ) . 
Our in vivo 3C assay does not allow us to draw equally firm conclusions , but nevertheless suggests that if an interaction does exist , it is likely to be weak , disrupted by our assays , or dependent on phage proteins not present in our 3C experiment . 
If an interaction does not exist , then the phage enzyme clearly has different properties from its mammalian counterparts , with which it shares only minimal structural homology [ 38 ] . 
But , then , Nature must find other ways of immobilizing the phage enzyme , or otherwise preventing the entanglement of nascent transcripts about their templates [ 11,39 ] . 
Materials and Methods
Templates
Template DNA was created by PCR from pLSG407 [ 40 ] unless otherwise indicated . 
KRF3/28 was the product of a PCR using primers KRF3 and KRF28 . 
The ` 452-bp template ' ( created using KFR3/28 as a template ) was the product of primers KRF28 and KRF32 , and contained a 59 biotin , followed by a BamHI site , a T7 promoter , and a 382-bp C-less cassette followed by 16 bp of Ccontaining DNA . 
The ` 290-bp template ' contained a T7 promoter followed by a 243-bp C-less cassette and 12 bp of C-containing DNA , and was the product of primers KRF36 and KRF37 . 
The ` 70-bp template ' was created using the oligonucleotide template KRF47 in combination with the primers KRF42 and KRF45 , and contained a T7 promoter followed by a 23-bp C-less cassette and 12 bp of C-containing DNA . 
Template DNA was purified using a 
Minelute PCR purification kit (Qiagen).
Labeling of DNA with fluors
The fluorescently-labelled 70-bp DNA template was prepared in the same manner as the unlabeled template , except that the primer KRF43 was replaced by the fluorescently-labeled primer KRF45 ( see Table S1 for primer sequence ) . 
KRF45 contained an amine-labeled dT residue near its 59 end , and was labeled using succinimidyl esters of Cy3B ( GE Healthcare ) or Atto647 ( Atto-Tec ) following the manufacturer 's instructions . 
One hundred micrograms of KRF45 was dissolved in 100 mL of H2O and extracted three times with an equal volume of chloroform . 
After the addition of 10 mL 3 M sodium chloride and 250 mL ethanol , the oligonucleotide was incubated at 220uC for 30 min , and then centrifuged at 12,000 * g for 30 min at 4uC . 
The pellet was allowed to dry , resuspended in 75 mL of 0.1 M sodium borate ( pH 8.5 ) , and frozen in 25 mL aliquots . 
A 50 nmol aliquot of succinimidyl ester was then resuspended in 5 mL DMSO , mixed with a 25 mL aliquot of KRF45 , and left overnight ( in darkness ) at 25uC . 
Labeled oligonucleotides were purified away from unconjugated fluorophore by ethanol precipitation , followed by one wash with 70 % ethanol . 
Comparing the absorbance of the oligonucleotide at 260 nm ( using e = 193,750 M 21 21 260 cm ) with its absorbance at 563 nm ( for Cy3b ; using e563 = 130,000 M cm , CF260 = 0.08 ) or 650 nm ( for Atto647N ; 21 21 e = 150,000 M 21 21 650 cm , CF260 = 0.06 ) showed that 90 -- 100 % of oligonucleotides were labeled . 
Denaturing urea-PAGE followed by visualization of the unstained gel with a FLA5000 imager showed that .90 % of the dye migrated with the purified oligonucleotide . 
The transcription buffer used in this experiment was either low-salt buffer ( LS1 ; 40 mM Tris-acetate pH 7.6 , 10 mM potassium chloride , 15 mM magnesium acetate , 5 mM dithiothreitol , 0.1 mg/mL N,N-dimethylated casein , 0.05 % Tween 20 , 0.4 U / mL RNase inhibitor , Roche ) or the more physiological potassium-glutamate buffer ( KGB ; 40 mM Tris-acetate pH 7.6 , 100 mM potassium glutamate , 15 mM magnesium acetate , 5 mM dithiothreitol , 0.1 mg/mL N,N-dimethylated casein , 0.4 U/mL RNase inhibitor ; [ 41 ] ) . 
The buffer LS1 was used because a study of the effect of buffer composition on T7 RNAP activity found this formulation to be optimal [ 24 ] . 
The buffer KGB was used because it is thought to mimic the cellular milieu [ 41 ] . 
The blocking agent in KGB was changed from bovine serum albumin ( BSA ) to casein because the latter yielded slightly higher T7 RNAP activity [ 24 ] . 
The experiment was performed at 25uC ( when LS1 was used ) or 37uC ( when KGB was used ) . 
A 60 mL transcription reaction contained transcription buffer plus 4 pmol His6-tagged T7 RNA polymerase , 0.6 pmol biotinylated 452-bp template , 0.6 pmol 290-bp template , and 0.2 pmol 800-bp control DNA . 
Two samples ( 2 mL each ) were taken , and immediately added to 10 mL ice-cold 16 TBE loading dye ( 89 mM Tris-borate , 89 mM boric acid , 2 mM EDTA , 0.05 % bromophenol blue ) . 
Separately , 30 mL of M270 magnetic 8 streptavidin beads ( 6.7610 beads per mL ; Invitrogen ) were washed twice in 200 mL transcription buffer , and then resuspended in the remaining 56 mL of the transcription reaction . 
After incubation for 20 min ( with mixing after 10 min ) , ATP , UTP , and GTP were added to a final concentration of 0.5 mM . 
Then , after 30 s , beads were pelleted with the aid of a magnet , and the supernatant removed . 
After removing a 2 mL sample ( and addition to TBE loading dye as above ) , supernatants were heated to 65uC for 10 min , and treated with 10 U RNase I ( Promega ) for 10 min at 37uC . 
The pellet was resuspended in water , then 106 LS1 was added to a final concentration of 16 , followed by the addition of 10 U/mL RNase I and 10 U BamHI ( assuring the initial ,60 mL volume was conserved ) . 
After 20 min at 37uC , beads were pelleted , the supernatant heated to 65uC for 10 min , and 2 mL samples collected ( and added to TBE loading dye as above ) . 
Fluorescence correlation spectroscopy
Transcription reactions ( performed in LS1 ) were initiated by addition of ATP , UTP , and GTP to 0.5 mM , and incubated for 30 s at 25uC before being pipetted onto a cleaned coverslip at 25uC . 
Fluorescence correlation spectroscopy was performed as described [ 42 ] . 
Time traces were acquired for 10 s using a SPQR ¬ 
14 avalanche photodiode ( Perkin Elmer ) , and autocorrelation functions were produced in real-time using a Flex02-02D correlation card ( Correlator.com ) . 
As our setup has a large pinhole , and therefore an elongated confocal spot ( longitudinal radius , wz . . 
wxy , the axial radius ) , translational diffusion times ( tD ) were extracted from autocorrelation curves by fitting to a two-dimensional single-species model , 1 t { 1 G ( t ) ~ ( 1z ) ( equation 1 ; [ 43 ] ) , where t is the delay time , N tD G ( t ) is the autocorrelation function , and N is the mean number of fluorescent molecules in the observation volume over the measurement . 
Experimentally acquired FCS curves were fit very well by this model ( e.g. , Fig. 2B and Fig . 
S4 ) . 
Although the molecules we analyze diffuse in three dimensions , the 3D model , 1 t t G ( t ) ~ ( 1z ) ( 1z ) ( where A = wz/wxy ; equation { 1 { 0:5 N t 2 D A tD 2 ; [ 28 ] ) , simplifies to the two-dimensional model ( equation 1 ) in the case of an elongated confocal spot [ 44 ] . 
To ensure that the 2D model was appropriate for modeling our data , we fit our Rhodamine 6G autocorrelation curves with both the 2D and 3D models . 
Fitting the data with the 3D model did not significantly change the values we obtained for either tD or N , however A could not be fit with reasonable confidence intervals ; changing the value of A therefore did not substantially affect the goodness of fit , a behavior consistent with confocal volumes where wz . . 
wxy . 
To ensure that our choice of model did not change the conclusions of our FCS work , we re-fit all of our FCS curves ( i.e. , all the data in Fig. 2A ) using the 3D model and setting A = 7 , a common value for single-photon excitation setups ; doing so increased all tD values by a small amount ( ,3 -- 5 % ) , with the difference between any two tD values changing by not more than 2 % . 
Two-species curves were calculated using the model 1 t { 1 G ( t ) ~ ( N1D1 ( t ) zN2D2 ( t ) ) , where Di ~ ( 1z ) , and N2 tDi N1 and N2 are the mean number of fluorescent molecules of species 1 and 2 , respectively , in the observation volume ( equation 3 ; [ 28 ] ) . 
Curve fitting was performed in MATLAB ( Mathworks ) . 
These models were also used to calculate the curves in Figure 2B . 
Fluorescence fluctuations were unlikely to be the result of dyespecific or photoinduced-photophysics , as the fitted N and tD of the fluorescently-labelled 70-bp template were unchanged when Atto647N was substituted for Cy3B , or when laser power was increased 10-fold ( data not shown ) . 
In order to convert diffusion times ( which depend on the size of the observation volume generated by the FCS setup ) into diffusion coefficients ( which are physical constants ) , we calculated the radius of the observation volume , v , using t 2 D ~ v = 4D ( equation 4 ; [ 28 ] ) . 
Measuring a diffusion time of 0.3860.1 ms ( fitting to equation 1 ) for the fluorescent standard rhodamine 6G ( D = 4.14 ? 
10 cm / s ; 26 2 [ 45 ] ) allowed us to calculate v = 7806100 nm . 
This observation volume is slightly larger than usual in order to maximize the number of photons captured from fluorophores during singlemolecule FRET experiments carried out on the setup ; however , this does not affect our ability to measure diffusion times . 
Chromosome conformation capture
This protocol -- modified from the original [ 30 ] for use in bacteria -- was generously provided by Mark Umbarger ( Harvard ; [ 46 ] ) . 
The E. coli strain KF22-1 was grown overnight to saturation in LB +50 mg/mL kanamycin , diluted by 1:250 into flasks containing 25 mL of the same media ( preheated to 37uC ) , and incubated at 37uC with shaking . 
After 30 min , arabinose was added to 0.4 % , or glucose was added to 0.2 % . 
When the cultures reached an OD600 of 0.4 , sodium phosphate ( pH 7.6 ) and formaldehyde were added to final concentrations of 10 mM and 1 % respectively ( except for non-crosslinked controls ) . 
After 20-min incubation at 37uC and 20-min incubation in an ice bath ( both with light shaking ) the formaldehyde reactions were quenched by addition of glycine to 0.125 M , and incubated for 5 min at 25uC . 
All cultures were then spun down at 5000 * g for 10 min , washed once with 50 mL ice-cold Tris-buffered saline ( 20 mM Tris-HCl pH 7.5 , 150 mM NaCl ) , pelleted , and stored at 280uC . 
The pellets were then resuspended in 1 mL TE buffer ( 10 mM Tris , 1 mM EDTA , pH 8 ) , and minor adjustments were made to assure that the OD600 of all samples was equal . 
For each pellet , 60 kU of Ready-Lyse Lysozyme ( Epicentre ) was added , and the mixture incubated at 25uC for 15 min with occasional gentle pipetting to resuspend cells . 
SDS was then added to a final concentration of 0.5 % and cells were allowed to incubate for 30 min . 
Five microlitres of solubilized chromatin ( ,100 ng DNA ) were mixed into a 50 mL volume containing 16 restriction buffer # 3 
( New England Biolabs ) and 1 % Triton X-100 , and incubated for 20 min to allow the Triton to neutralize the SDS . 
Fifty units of BglII ( New England Biolabs ) were added , and the chromatin digested for 2.5 h at 37uC with light shaking . 
One additional sample served as a no-restriction enzyme control . 
The reaction was then halted by addition of SDS to 1 % . 
In order to form intra-molecular ligation products , 60 mL digested chromatin was added to 760 mL ` ligation mix ' containing 16 T4 ligase buffer , 1 mM ATP , 25 mg/mL BSA , 1 % Triton X-100 , and 2.4 kU/mL T4 DNA ligase . 
One additional sample served as a ` no ligase ' control . 
Ligase mixtures were then incubated for 16uC for 1 hr . 
The reaction was halted by the addition of EDTA to 10 mM , and incubated overnight with 50 mg of proteinase K at 65uC . 
Four hundred microlitres of the DNA solution was then extracted twice with 400 mL of 25:25:1 phenol : chloroform : isoamyl alcohol . 
Glycogen was added to a final concentration of 50 mg/mL . 
Ice-cold sodium acetate and ethanol were then added to final concentrations of 0.75 M and 70 % ( v/v ) respectively . 
The DNA-glycogen mixture was incubated at 280uC for 3 h , and then spun down at 20,000 * g at 4uC for 20 min . 
The pellet was then washed with 1 mL 70 % ( v/v ) ethanol ( 25uC ) , air dried , and resuspended in 12 mL distilled , deionized , H2O . 
PCR was performed using FlexiTaq DNA polymerase ( Pro-mega ) and 16 reaction buffer , 1.75 mM MgCl2 , 0.2 mM dNTPs , 0.4 mM primers and 2 % DMSO on a thermocycler using the following program : ( i ) 95uC for 1 min , ( ii ) 95uC for 1 min , ( iii ) 65uC for 45 s , ( iv ) 72uC for 2 min , ( v ) repeat steps ii -- iv 35 times , and ( vi ) 72uC for 6 min . 
Ligations between restriction fragments 1 ( T7 gene 10 ) and 8 ( control DNA fragment ) were amplified using primers KF101to8BglIIfw and KF101to8BglIIrv ; these primers were designed to produce a fragment of 243 bp ( this corresponded to ligation product a : b in Fig. 4A ; all primer sequences can be found in Table S1 ) . 
Ligations between restriction fragments 1 ( T7 gene 10 ) and 16 ( pT7-Ypet ) were amplified using primers KF101to16B-glIIfw and KF101to16BglIIrv ; these primers were designed to produce a fragment of 217 bp ( this corresponded to ligation product a : c in Fig. 4A ) . 
We queried the inversion and ligation of two adjacent fragments of E. coli genomic DNA by PCR using primers 3CposconA and 3CposconB ; these primers were designed to produce a fragment of 443 bp ( this corresponded to ligation product d : e in Fig. 4A ) . 
The identity of all PCR products was confirmed by measuring the size of the products , and by digesting these products with BglII ( data not shown ) . 
We quantified the amount of ligation products produced in our 3C reactions using PCR , following well established protocols [ 47 ] . 
We began by optimizing PCR conditions ( i.e. , amount of 3C template per reaction , and number of PCR cycles ) to ensure that the amount of PCR product produced was linearly related to the amount of ligation product initially present in the PCR reactions . 
This was accomplished by performing PCR reactions containing serial dilutions of the 3C template , subjecting the PCR reactions to gel electrophoresis ( on a TBE-2 % agarose gel ) , staining the gels with SYBR green I , and measuring the intensities of the bands corresponding to the amplification products ( using AIDA image analysis software ) . 
We found that , for all the ligation products we examined , 36 PCR cycles on 30 ng of our 3C template resulted in bands with an intensity that was proportional to the amount of ligation product in the initial PCR reactions ( e.g , see Fig. 4B lanes 1 , 5 , and 6 ) . 
Using these conditions , we then conducted PCR on all experimental samples in triplicate . 
For each primer pair , controls containing 15 ng and 60 ng ` + T7 ' 3C template ( i.e. , 0.56 and 26 the normal amount ) were also included to ensure that the intensity of the bands produced on our gels was linearly related to the amount of ligation products in the PCR reactions ( these controls are found in Fig. 4B , lanes 1 , 5 and 6 ) . 
Only samples run on the same gel were directly compared . 
The goal of the experiment was to determine whether the interaction frequency of the transgenes PT7-gene10 and PT7-YFP , ( X ) , in the presence of T7 RNAP , XzT7 , was greater than the interaction frequency of these two genes in the absence of T7 RNAP , X { T7 . 
In other words , the goal was to determine whether XzT7 = X { T7 was greater than 1 . 
The relationship between interaction frequencies ( which occur in the cell ) and ligation frequencies ( which are present in a 3C template sample ) is given by ( LzT7 = LCzT7 ) X = X ~ ( equation 5 ) , where L and zT7 { T7 zT7 ( L { T7 = LC { T7 ) L { T7 are the ligation frequencies of the transgenes in the presence and absence of T7 RNAP , while LCzT7 and LC { T7 are the ligation frequencies of two control restriction fragments that should interact at the same rate regardless of whether or not the transgenes are transcribed by T7 RNAP ( these two control ligation products were amplified by primers a : b or d : e ; Fig. 4A ) . 
This equation states that directly comparing ligation frequencies between different 3C samples is possible only after differences in the efficiency of the 3C protocol between samples are controlled for . 
If we assume that the intensity of the band produced by each amplified ligation product is proportional to the original amount of ligation product in the 3C template ( we do , indeed show that this is the case , see above , and Fig. 4B lanes 1 , 5 , and 6 ) , then the intensity of the band seen in the gel , I , is related to the amount of ligation product in the PCR reaction , L , by L ~ a : I , where a is the efficiency of the relevant primer pair . 
Then ( aT7IzT7 = aCICzT7 ) ( IzT7 = ICzT7 ) XzT7 = X { T7 ~ ~ ( equation ( aT7I { T7 = aCIC { T7 ) ( I { T7 = IC { T7 ) 6 ) . 
This equation reveals that because the experiment is ultimately interested in a change in a single interaction frequency , primer efficiencies cancel out , and have no effect on the final result . 
It also gives the expressions that must be measured in order to determine whether the interaction frequency of the two transgenes changes in the presence of T7 RNAP . 
The values of ( IzT7 = ICzT7 ) and ( I { T7 = IC { T7 ) are given by the ` test gene contact frequencies ' in Fig. 4B lanes 1 and 2 . 
Because these values are virtually identical , XzT7 = X { T7 is ,1 . 
This result indicates that the interaction frequency of the transgenes is not changed by the presence of T7 RNAP . 
To test the efficiency of restriction nuclease digestion , PCR primers BglIIconfw and BglIIconrv were chosen to amplify a 285 bp fragment of genomic DNA containing a BglII site at its centre . 
To quantify total DNA , PCR primers rpoZampfw and rpoZamprv were chosen to amplify a 292 bp genomic fragment that did not contain a BglII site . 
Restriction digestion efficiency was determined by comparing the ratios of the BglIIconfw/rv fragment : rpoZampfw/rv fragments in the presence and absence of restriction digestion . 
Supporting Information
based assays . 
A. Diagrams of DNA fragments ( i ) 800-bp promoter-less control fragment . 
( ii ) 452-bp template . 
( iii ) 290-bp template . 
( iv ) 70-bp template . 
Numbers indicate the position of elements ( in bp ) relative to the 59 ends of the templates . 
B. Transcripts produced by T7 RNAP . 
The templates in ( A ) were transcribed in reactions containing 16 KGB , 100 nM template , 200 nM RNAP , and 0.5 mM ATP+GTP + [ 32P ] UTP ( 0.25 mCi / mL ) in the presence or absence of 0.5 mM CTP . 
After 10 min , the resulting RNA was separated by denaturing urea-PAGE , and visualized using a phosphoimager screen ( Molecular Dynamics ) and a FLA5000 imager ( Fuji ) . 
( i ) Transcripts produced by all three templates . 
( ii ) A second gel better resolving the transcripts produced using the 452-bp template ( below ) . 
The shorter products produced in reactions lacking CTP indicate that RNAPs transcribe the C-less cassettes but halt at the first C residue . 
( TIF ) reaction ( in buffer LS1 ) lacking NTPs containing 50 nM T7 RNAP and 8 nM of the 452-bp template ( encoding a T7 promoter , a 382-bp C-less cassette , and a C-containing 39 end ) was prepared , and sampled under sequentially-applied conditions . 
These samples were separated using a native 1.5 % agarose gel , and stained with SYBR green I . 
In the absence of NTPs , the templates are not stably bound by RNAPs , and thus migrate as free DNA ( lane 1 ) . 
Adding ATP+UTP+GTP ( to 0.5 mM ) causes 
RNAPs to initiate and halt at the end of the C-less cassette . 
The templates are now stably bound by RNAPs and their transcripts , and so migrate more slowly ( lane 2 ) . 
Adding CTP ( to 0.5 mM ) allows RNAPs to ` run-off ' and vacate most templates , which migrate once again as free DNA ( lane 3 ) . 
DNase treatment shows that RNA makes only a minor contribution to the observed fluorescence ( lane 4 ) , while additional RNase treatment removes all nucleic acid ( lane 5 ) . 
B . 
The fraction of template occupied by T7 RNAP in ( B ) quantified using AIDA image-analysis software ( Raytest ) . 
For each condition , the amount of occupied template was calculated by subtracting the amount of freely-migrating DNA ( as judged by band intensity ) from the total amount of DNA ( found in lane 1 ) . 
Repeating the experiment in the buffer KGB instead of LS1 yielded similar results ( data not shown ) . 
( TIF ) duced during the ` pulldown ' assay . 
A transcription reaction ( in KGB ) containing 0.1 mM biotinylated 452-bp template , 0.1 mM 290-bp template , and 0.3 mM T7 RNAP was initiated by the addition of ATP+GTP + [ 32P ] UTP ( 0.25 mCi/mL ) to 0.5 mM in the presence or absence of beads ( 4.56108 beads/mL ) . 
After 30 s , reactions were halted by the addition of formamide to 80 % ( v/v ) , and subjected to denaturing urea-PAGE . 
Total [ P ] RNA was 32 then visualized using a phosphoimager screen ( Molecular Dynamics ) and a FLA5000 imager ( Fuji ) . 
B. Quantitation of the 32 P incorporated into the transcripts in ( A ) . 
Initiation rates on the 452-bp and 290-bp templates can be inferred from the intensities of the corresponding transcripts ( which measured 382 bp and 243 bp , respectively ) . 
When transcript length is accounted for , we see that RNAPs initiated on the 452-bp template at ,0.76 the rate at which they initiated on 290-bp templates . 
We conclude that when the majority of 290-bp templates are occupied , a similar fraction of the 452-bp templates will also be occupied . 
species model . 
( i ) Representative autocorrelation curve ( blue , upper panel ) recorded using FCS in the experiment of Fig. 2Aiv . 
A reaction containing 1.75 mM T7 RNAP , 2 nM labeled 70-bp template , and 0.54 mM unlabeled 452-bp template , was initiated by the addition of ATP+UTP+GTP . 
After RNAPs had halted at the first C residues ( 30 s ) , the autocorrelation function of the labeled templates was determined by FCS . 
( ii ) A fit of the autocorrelation function produced in ( i ) using a two-dimensional one-species model ( red , upper panel ; equation 1 ) , and yielding a diffusion time of 4.1 ms. Residuals ( red , lower panel ) are minor , suggesting that the model used to fit the curve is well-suited to the sample ( see Materials and methods ) . 
( TIF )