Transcriptional regulation controls the ability of RNA polymerase to initiate the transcription of a gene into mRNA


1


. It is crucial in the life of all organisms, orchestrating the expression of genes in response to environmental cues and cellular states


<sup>2

,

3</sup>


. Transcriptional regulation is mediated by transcription factors (TFs), proteins that bind DNA near a gene at short DNA words known as transcription factor binding sites (TFBSs). The binding of a TF to its binding site on DNA can either hinder or facilitate transcription initiation by RNA polymerase


<sup>1

,

4

,

5</sup>


. The stronger the binding of a TF to its TFBS is, the more strongly the TF can regulate a nearby gene


<sup>6

–

9</sup>


. In bacteria, TFs operate within a hierarchically organized gene regulatory network. At the bottom of this hierarchy are

local

TFs that regulate the expression of one or few genes and modulate specific biological processes


<sup>10

–

12</sup>


. At the top are

global

TFs that may regulate hundreds of genes and control many cellular functions


<sup>11

,

13

–

15</sup>


. Consistent with their broader role, global regulators are typically more highly expressed and bind to a broader range of TFBSs than local regulators


¹²


.

A genotype-phenotype map is a conceptual analog of a physical landscape, where each location corresponds to one genotype in sequence space, and is associated with a quantitative phenotype. If the phenotype is related to gene expression, the map is also called a regulatory landscape


<sup>6

,

16

–

19</sup>


. Another special case of such a map is a fitness landscape or adaptive landscape, in which the phenotype is fitness and is interpreted as an elevation


<sup>20

,

21</sup>


. Over the last decade, genotype–phenotype (GP) maps and fitness landscapes have become central tools for understanding how molecular systems evolve under mutation and selection


<sup>22

–

25</sup>


. Such maps and landscapes have been experimentally studied for DNA


<sup>6

,

8

,

18

,

19

,

26

,

27</sup>


, protein


<sup>28

–

32</sup>


and RNA


<sup>33

–

35</sup>


molecules, revealing key topographical properties that shape evolutionary outcomes, including epistasis


<sup>24

,

36</sup>


—the non-additive effects of multiple mutations on phenotype—landscape ruggedness, reflected in the number and distribution of fitness peaks, and constraints on adaptive evolution. For example, one large-scale study adopted CRISPR-Cas9 technology to measure the fitness of more than 200,000

E. coli

genotypes that encode variants of the bacterial antibiotic resistance gene dihydrofolate reductase (DHFR). It showed that this landscape is highly rugged (multi-peaked)


³¹


.

For transcription factor binding sites, most pertinent large-scale studies are based on in vitro binding assays, such as protein-binding microarrays (PBMs), and they focus predominantly on eukaryotic transcription factors


⁶


. While these studies have been instrumental in characterizing transcription factor binding preferences, they typically do not measure regulatory output in a native cellular context. In contrast, comprehensive in vivo data for bacterial TFBSs remain extremely rare. To our knowledge, only two high-resolution in vivo landscapes have been previously mapped for bacterial regulators, those of the local regulators TetR


¹⁸


and LacI


²⁷


. As a result, it remains unclear whether principles inferred from protein landscapes, eukaryotic TFBSs, or in vitro binding assays generalize to transcriptional regulation in bacteria, particularly for global regulators


¹¹


that integrate multiple physiological signals.

Both TFs and their TFBSs evolve, but TFBSs evolve more rapidly. The reason is that a mutation in a TF can affect the expression of many genes, whereas a mutation in a TFBS may affect only one gene and is thus less likely to be deleterious


<sup>13

,

37

–

39</sup>


. A special case of TFBS evolution is the evolution of a strong TFBS from a DNA word with weak or no regulatory activity. Such

de novo

evolution of TFBSs can create new regulatory interactions and change the structure of gene regulatory networks


<sup>40

–

43</sup>


. Unfortunately, we know little about the ability of Darwinian evolution to create TFBSs de novo. A strong TFBS may have to arise from a non-binding site through an evolutionary path of multiple mutational steps. Darwinian evolution can favor this process only if strong binding is adaptive and if each mutational step in a path increases binding strength, i.e., if the path is

accessible

to Darwinian evolution. To find whether such paths exist may require the analysis of multiple paths. This is challenging because sequence space contains an astronomical number of potential TFBSs for any one TF. The number of evolutionary paths to strong transcriptional regulation is thus also astronomical. For each such path, the strength of each TFBS along the path has to be measured experimentally


<sup>44

–

47</sup>


.

In principle, one could attempt to construct such landscapes in silico using commonly employed models of TF–DNA interactions, such as position weight matrices (PWMs)


<sup>48

–

50</sup>


. However, PWMs are derived from a limited set of naturally occurring binding sites and primarily reflect sequence conservation rather than quantitative regulatory output. Moreover, PWMs assume independent and additive contributions of individual nucleotide positions to DNA binding


<sup>48

–

50</sup>


. They therefore cannot capture the influences of epistatic interactions between positions, which can dramatically affect landscape topography and evolutionary accessibility


²⁴


. Lastly, PWMs do not account for important biological effects that modulate gene regulation such as DNA shape


<sup>51

,

52</sup>


, cooperative interactions


<sup>53

,

54</sup>


, and chromosomal context


<sup>55

,

56</sup>


. Thus, experiments are necessary not only to obtain quantitative measurements of gene regulatory activity, but also to refine and inform PWM-based models using thousands of experimentally characterized sequences.

Building on our previous work on a local TF


¹⁸


, we address this challenge for three global regulators in

Escherichia coli

by performing three independent and massively parallel experiments


⁴⁴


for each TF. The experiments use a synthetic biology platform to quantify how strongly each of more than 30,000 binding sites for a TF can regulate the expression of a nearby reporter gene.

The first TF we study is the cAMP receptor protein (CRP). In the absence of glucose, CRP modulates the expression of genes mostly involved in carbon metabolism. It allows

E. coli

to efficiently switch between sources of carbon and energy


<sup>57

–

60</sup>


. The second TF is the factor for inversion stimulation (Fis), which helps to regulate the expression of genes involved in growth phase transitions. It also modulates the supercoiling of DNA


<sup>61

–

63</sup>


and influences DNA replication, recombination, and repair


<sup>61

–

63</sup>


. The third factor is the integration host factor (IHF). IHF regulates genes involved in stress responses and stationary phase survival


<sup>64

,

65</sup>


. Similar to Fis, it is involved in DNA compaction, replication, and recombination, but also in the assembly of complex nucleoprotein structures


<sup>64

,

65</sup>


. We chose these factors because they are the most global regulators in

E. coli

, and they are diverse, belonging to different protein families.

We use our experimental data for each TF’s binding sites to map the relationship between binding site genotype and gene expression. Our first aim is to characterize the resulting regulatory landscapes for global bacterial regulators, and to find out whether these landscapes are different or similar. When strong regulation of a gene is adaptive, a regulatory landscape becomes a fitness landscape


<sup>20

,

21

,

25

,

66</sup>


. Our second aim is to understand how populations would evolve on each of our three landscapes when they are viewed as fitness landscapes. Specifically, we study how evolving populations would traverse each landscape through individual mutational steps that change a TFBS its ability to regulate a gene via its cognate TF. A peak in such a landscape is a TFBS conveying stronger regulation than all neighboring TFBSs in sequence space. If such a landscape is rugged (has multiple peaks), natural selection alone may not enable a population to discover the highest peaks, i.e., the strongest TFBSs. The reason is that the peaks may be separated by valleys of low regulation strength that cannot be traversed by natural selection alone


<sup>67

–

69




25</sup>


. In other words, high peaks may not be easily accessible through Darwinian evolution – they may be reachable by few or no evolutionary paths of single DNA mutations in which each mutational step increases TFBS strength


⁶


. One factor that can reduce peak accessibility is epistasis, which can reduce the predictability of evolutionary trajectories towards a peak


<sup>24

,

36</sup>


.

To accomplish both aims, we first studied the topography of the three landscapes, and then simulated the dynamics of evolving populations on them. All three landscapes are highly rugged. They contain more than 2,000 peaks that are scattered through genotype space and are rife with epistatic interactions, in striking contrast to the comparatively smooth TFBS landscapes described for eukaryotic systems


⁶


. Despite these features, evolving populations can reach the strongest TFBSs in all three landscapes. Which of several high peak is reached is contingent on chance events during adaptive evolution.

Results

Landscape mapping

We constructed a modular plasmid system based on the common backbone plasmid pCAW-Sort-Seq-V2


¹⁸


(

Supplementary Table S1

;

Supplementary Figures S1

–

S2

;

Supplementary Methods 2–3

). This backbone contains all shared regulatory and reporter elements required for fluorescence-based measurements, but it encodes neither a TF nor its binding site(s) for any of the regulators studied here. From this backbone, we generated three TF-specific plasmid derivatives. Each of these plasmids encodes one of the global transcription factors CRP, Fis, or IHF under inducible control (hereafter pCAW-Sort-Seq-V2-CRP, pCAW-Sort-Seq-V2-Fis, and pCAW-Sort-Seq-V2-IHF;

Supplementary Figure S2

). In each of the TF-specific plasmids, a TFBS insertion site is positioned immediately upstream of the

gfp

reporter gene, such that TF binding represses the transcription of

gfp

. Consequently, stronger TF–DNA binding results in lower GFP expression, which enables a quantitative readout of a binding site’s regulation strength.

For each TF-specific plasmid, we constructed three kinds of variants. The first carries a wild-type (WT) TFBS for the corresponding TF upstream of

gfp

. It serves as a reference conferring wild-type regulation of the reporter gene. The second contains a complete TFBS library, in which we randomized the eight most information-rich base-pair positions of the respective binding site (

Supplementary Methods 4

;

Supplementary Table S3

), as determined from alignments of experimentally characterized binding sites curated in RegulonDB


⁷⁰


. Each library comprised 4⁸ = 65,536 unique TFBS sequences, and we constructed three independent biological replicate plasmid libraries from them per TF (

Supplementary Methods 4–5

). The third variant lacks a promoter upstream of

gfp

and serves as a negative control that allows us to quantify cellular autofluorescence during fluorescence measurements (

Supplementary Table S1

).

We introduced each TF-specific plasmid into an

E. coli

host strain in which the corresponding chromosomal TF gene had been deleted (

Δcrp

,

Δfis

, or

Δihf

;

Supplementary Table S2

). This genetic background ensures that the focal transcription factor is expressed exclusively from the plasmid. Although the mutant strains grow more slowly than the WT, they reached similar cell densities during late exponential or early stationary phase, the growth phase at which we performed all measurements (

Supplementary Figure S3

). TF expression is controlled by a tetracycline-inducible promoter and can be precisely tuned using anhydrotetracycline (aTc), allowing us to regulate TF abundance independently of growth conditions and to isolate the effects of TF–DNA binding on transcriptional regulation (

Figure 1a

).

We then mapped these TFBS genotypes to their respective regulatory phenotypes using a well-established technique known as sort-seq


<sup>16

,

44

,

71

–

73</sup>


(

Figure 1b

), which combines fluorescence-activated cell sorting (FACS) with high-throughput sequencing (

Figure 1c

). In sort-seq, one first sorts cells into multiple fluorescence “bins” depending on the level of GFP expression. A cell’s GFP fluorescence serves as a proxy for GFP expression levels


⁷⁴


and quantifies how strongly the TFBS library member in this cell can regulate GFP expression (

Methods

,

Supplementary Methods 5

). For each TFBS genotype, we quantified regulation strength (S) as a weighted average of its sequencing counts across the different fluorescence bins, yielding a single continuous measure of regulatory activity (see

Methods

).

The results of our sort-seq experiments are three maps – one for each transcription factor – from each of more than 30,000 genotypes (TFBS variants) to regulatory phenotypes (regulation strength). One can view each map as a regulatory landscape


<sup>8

,

9

,

16

,

18

,

75

–

79</sup>


(

Figure 1d

) that becomes a fitness landscape whenever strong regulation entails high fitness


<sup>6

,

25</sup>


. For the purpose of analyzing the landscape quantitatively, we represent it as a network of genotypes (TFBS variants). Each node in this network corresponds to a genotype. Edges link neighboring genotypes, which differ in a single nucleotide (

Figure 1d

).

Landscapes exhibit diverse regulation strengths and distribution breadths

To evaluate the ability of our library to regulate gene expression, we first measured the distribution of GFP fluorescence intensities across the bins in two conditions, i.e., in the presence or absence of the TF (with or without the atc inducer,

Supplementary figures S4

-

S6

). In the presence of the TF, GFP expression was lower on average and showed a broader distribution than in the absence of the TF (

Supplementary figures S4

-

S6

). This indicates that the TFBSs in each library can indeed downregulate GFP expression, but some library variants convey stronger regulation than others, hence the broader fluorescence distribution.

We then pooled barcoded DNA sequences extracted from cells in each bin and biological replicate. We sequenced at least 250 unique TFBS genotypes from each bin, with an average of 3992.3±4293.9 unique sequences per bin for the three TFs (as detailed in

Methods

,

Supplementary Methods 7

, and

Supplementary Figures S4

-

S6

). The resulting sequences covered 95%, 90%, and 93% of the total library sizes (N = 65,536 genotypes) for CRP, Fis and IHF, respectively. To ensure the reliability of our data, we excluded sequences with low read coverage and sequences that were not present in all triplicates (

Methods, Supplementary Methods 7.3

). This quality filtering step resulted in library sizes of 31,975 genotypes for CRP (49% of all 4

⁸

genotypes), 43,222 genotypes for Fis (66%), and 41,325 genotypes for IHF (63%). The correlation in read counts for each variant across replicates was very high for this quality-filtered data (Pearson’s R= 0.98-0.99) (

Supplementary Figures S7

-

S9

). We used this data for all further analyses.

The fluorescence and sequence data from each bin allowed us to quantify the variant’s ability to regulate gene expression for each TFBS library variant. We refer to the resulting metric as the

regulation strength S

conveyed by the variant (

Methods

,

Supplementary Methods 7.2

,

Figure 1d

). It ranges between S=0 (no regulation) to S=1 (strongest regulation among all variants in the library). Although our experiments directly quantify expression regulation, and not the affinity or binding strength of a TF to a TFBS variant, we also use

binding strength

as a proxy for regulation strength, because TF-DNA binding is necessary for regulation


<sup>1

,

4

,

11

,

54

,

80

,

81</sup>


. To each of our three TFs, we also assigned a reference TFBS that is naturally occurring and conveys strong regulation by the TF, as proven by previous experimental work


<sup>60

,

82

–

84</sup>


. We refer to this TFBS as the wild-type (WT


<sup>82

,

85</sup>


WT


<sup>83

,

86</sup>


and WT


<sup>84

,

87</sup>


,

Supplementary Table S4

). We refer to TFBSs with regulation strengths below and above the wild-type (WT

_CRP

: S

_WT

=0.71, WT

_Fis

: S

_WT

=0.97, WT

_IHF

: S

_WT

=0.95) as weak and strong, respectively.

We observed a broad range of regulation strengths S for each TF landscape, with varying dispersions. The CRP landscape exhibited the lowest average regulation strength and a narrower distribution of S compared to Fis and IHF, which had similar distributions (mean±s.d., 0.37±0.1 for CRP, 0.57±0.13 for Fis, and 0.52±0.14 for IHF; see

Figure 2a

,

Supplementary Figure S10

). Next, we analyzed the strongly regulating TFBSs to identify nucleotides that may be particularly frequent, and thus potentially important for strong regulation (

Figure 2b

). We discovered a moderate association between the most frequent nucleotides (highlighted in yellow in

Figure 2b

) and the most informative nucleotides from the available position weight matrices (PWMs) for these TFs


⁷⁰


. (Pearson correlation coefficients: R=0.51, R=0.43, R=0.47 for Fis, CRP and IHF, respectively, with p-values smaller than 10

^-16

, rejecting the null hypothesis of an absence of association). This observation suggests that our logos capture different information compared to available PWMs. This is expected, because our approach allows us to filter sequences by regulation strength thresholds to construct our matrices, unlike the traditional method of aligning genomic TFBSs without considering their regulation strengths


<sup>48

,

49</sup>


.

To further validate the data from our sort-seq experiments, we isolated cells harboring 10 different TFBS variants from each of 13 bins of each library (i.e., 10×13=130 variants per library), and determined their regulation strength more directly by quantifying GFP expression with a microplate reader (

Methods

and

Supplementary Figure S11

). This comparison validates the sort seq approach by revealing a strong association of the two independent quantifications of regulation strength (Pearson’s R=-0.81 for CRP, R=-0.74 for Fis, and R=-0.73 for IHF). (

Methods

and

Supplementary Figure S11

).

All three regulatory landscapes are highly rugged

The study of our landscapes in a network framework (

Figure 1d

) can help to quantify different aspects of landscape topography (

Supplementary Table S5

). One of them is the ruggedness of each landscape. It can be quantified by the number of peaks


<sup>31

,

88

–

90</sup>


. In the network framework, a peak is a TFBS whose neighbors are all weaker regulators (with lower S) than the TFBS itself. We find that all three landscapes are highly rugged, with 2,154, 2,312, and 2,453 peaks for the CRP, Fis, and IHF landscapes, respectively (

Figure 2c

,

Supplementary Figure S12

,

Supplementary Table S5

). Not surprisingly, peak genotypes generally are stronger regulators than non-peak genotypes (

Figure 2c

). Only a small fraction of peak genotypes regulate expression more strongly than the wild-type (

Figure 2c

; 61 for CRP, 172 for Fis, and 199 for IHF). We refer to such peaks as high peaks and distinguish them from low peaks (S<S

_WT

).

The prevalence of sign epistasis (

Supplementary Table S5

) supports the notion that our landscapes are indeed rugged (see

Supplementary Methods 7.5

for further details on epistasis and its evolutionary consequences). Independent evidence for landscape ruggedness comes from comparing the ruggedness of our landscapes with that of a well-established theoretical model of uncorrelated random landscapes, in which each sequence is assigned a fitness at random from the same fitness distribution, and neighbors have uncorrelated fitness values


<sup>90

–

92</sup>


. Such landscapes are maximally rugged


<sup>90

–

92</sup>


. We created 10

³

uncorrelated random landscapes for each of our three TF landscapes by randomly shuffling the measured regulation strengths among all genotypes (

Supplementary Methods 7.6

). The number of peaks in our TF landscapes lies within 93%, 96%, and 98% percent of that of a maximally rugged random landscape, which, on average [mean±s.d.], has 2,308±133, 2,405±89 and 2,373±97 peaks for CRP, Fis and IHF, respectively. This analysis underlines that our landscapes are indeed highly rugged.

Because we use only quality-filtered genotype data, our landscapes lack regulatory information for about 40% of the 4

⁸

genotypes. While we cannot exclude that this undersampling of genotypes has led to systematic biases in our determination of landscape ruggedness, we note that the sampling of landscape genotypes by our experiments was not strongly biased with respect to regulation strength. Specifically, when we analyzed the relative connectivity of genotypes in our landscapes – the fraction of each genotype’s 24 possible neighbors for which our experiments yielded regulatory data – we found that it is only weakly correlated with regulation strength (R=-0.1, -0.1, 0.01 for the CRP, Fis, and IHF landscapes,

Figures S13

-

S15

). Similarly, the relative connectivity of peak genotypes is only weakly correlated with their regulation strength (R=-0.05, -0.04, 0.06 for the CRP, Fis, and IHF landscapes).

Landscape peaks are widely scattered in genotype space

In a rugged adaptive landscape, reaching high fitness peaks can be challenging, because such peaks are separated from other genotypes by valleys of low fitness that cannot be traversed by natural selection alone


²⁵


. If natural selection favors strong gene regulation, the ruggedness of our regulatory landscapes may thus present a challenge for adaptive evolution. To better understand the potential magnitude of this challenge, we next analyzed our landscapes’ topography in greater detail. We began by studying the distribution of peaks in genotype space. If a landscape’s highest peaks are widely scattered through genotype space, then they may be accessible via fitness-increasing evolutionary paths from diverse non-peak genotypes. This may facilitate adaptive evolution compared to a landscape where peaks are clustered in a small region of genotype space.

For each of our three landscapes, we determined the genetic distance between peaks, i.e., the minimum number of mutations needed to transition from one peak to another, regardless of their effect on regulation strength. We compared the distribution of these distances to the distribution of genetic distances for an equal number of randomly selected non-peak variants. In all three landscapes, the distances between peaks are almost indistinguishable from those of random genotypes, differing on average by fewer than 0.1 mutational steps. In other words, peaks are about as widely dispersed in each landscape as random genotypes (

Supplementary Figure S16

). This is the case for both low and high peaks (

Supplementary Figures S17

-

S18

). A principal component analysis further underscores this dispersion (

Supplementary Methods 7.7

,

Supplementary Figures S19

-

S21

).

Accessible paths to a peak are often not the shortest possible paths

Next, we focused on mutational paths to peaks that are evolutionarily accessible, i.e., a series of mutational steps where each step increases regulation strength. Specifically, we enumerated all accessible paths that exist from each non-peak genotype to each peak genotype for all three landscapes. We found that these paths are generally longer than the shortest genetic distance between a non-peak and the peak genotypes. Specifically, the mean length of accessible paths exceeded the shortest distance by 1.5, 1.8 and 1.8 steps for the CRP, Fis and IHF landscapes (

Supplementary Figure S22

). In other words, accessible mutational paths are often not the most direct paths.

The existence of indirect paths implies that some mutational steps are evolutionarily prohibited because they decrease regulation strength. The reason is closely linked to non-additive (epistatic) effects of two or more mutations on regulation strength. (see

Supplementary Methods 7.5

for further details on epistasis). More specifically, such inaccessible steps are a result of sign epistasis. In this kind of epistasis, a double mutant of a TFBS regulates expression more strongly than the TFBS itself, even though one or both constituent single mutants regulate expression more weakly than the TFBS


<sup>24

,

93

–

95</sup>


. Indeed, epistasis is prevalent in all three landscapes (

Table S5

). Specifically, we observe sign epistasis in 62%, 66%, and 65% percent of interactions between single mutant pairs in the CRP, Fis, and IHF landscapes.

The existence of accessible paths alone does not tell us how easily a high peak can be found through Darwinian evolution. The reason is that only a very small fraction of all paths to that peak may be accessible, and an evolving population may not find any one of those paths. To quantify the fraction of accessible paths, we determined the total number of paths from each non-peak genotype to each high fitness peak, and computed the fraction of those paths that are accessible.

Figure 3a

shows how this fraction depends on the number of mutational steps in a path. It behaves similarly for all three landscapes (

Figure 3a

). The majority of two-step paths (a fraction greater than 80%) are accessible in all three landscapes (

Figure 3a

), but the fraction of accessible paths dwindles rapidly with an increasing number of mutational steps. It reaches a minimum below 1% for all three landscapes at eight mutational steps (

Figure 3a

).

High peaks have large basins of attraction that share many TFBS genotypes

In the following analysis, we computed the basin of attraction of each peak, defined as the set of non-peak genotypes from which accessible paths to the peak exist


<sup>31

,

96

,

97</sup>


. In other words, a peak’s basin of attraction comprises all genotypes from which adaptive evolution can access the peak. The peaks in our landscapes have widely different basin sizes. They include peaks accessible from a mere fraction of genotypes to those accessible by a majority of genotypes (

Figure 3b

). Notably, high peaks generally had larger basins of attraction than low peaks (

Figure 3c

,

Supplementary Figure S23

). In addition, we found that many genotypes are part of multiple basins of attraction. Adaptive evolution may reach different high peaks starting from any such genotype. For all pairs of high peaks, we computed the pairwise overlap (intersection) of the basins of attraction, i.e., the fraction of genotypes that are part of both basins of attraction. (

Figure 3d

,

Supplementary Figures S24

-

S25

). The number of genotypes in this intersection varies widely, but the basins of attraction of high peaks generally share a substantial fraction of genotypes (mean shared genotypes: 48%, 42%, and 36% among all pairs of high peaks for the CRP, Fis, and IHF landscapes,

Figure 3d

)

Genetic drift facilitates and clonal interference reduces the attainment of high peaks

Taken together, our analyses thus far indicate that high peaks in the CRP, Fis, and IHF landscapes are more accessible than low peaks (

Figure 3b

), and their basins of attraction also share more genotypes (

Figure 3d

). However, the accessible evolutionary paths to high peaks are often indirect. In addition, the fraction of accessible paths to any one high peak dwindles rapidly with the distance from that peak (

Figure 3a

).

To understand how these topographical features jointly affect adaptive evolution, we simulated the evolutionary dynamics on our three landscapes under the assumption that natural selection would favor strong regulation and that fitness is proportional to regulation strength.

Because high peaks constitute only a small fraction of all peaks (2.8%, 0.4%, and 0.5% in the CRP, Fis, and IHF landscapes), we hypothesized that most evolving populations would reach only low peaks. To test this hypothesis, we took advantage of the fact that

E.coli

has a low genomic mutation rate of 2.2 × 10

⁻¹⁰

per base pair per generation


⁶⁷


, and that we study evolution only within an 8-nucleotide segment of a TFBS. In addition,

E. coli

has a large effective population size (>10

⁸

individuals


⁶⁷


), which means that genetic drift is weak and even minor differences in fitness are visible to natural selection


<sup>67

,

98</sup>


. These conditions imply that a population on our landscape would evolve in the well-studied strong-selection weak-mutation regime (SSWM)


<sup>16

,

99

–

101</sup>


. In this regime, a population is monomorphic most of the time, until a beneficial mutation arises, which usually sweeps rapidly to fixation. In other words, adaptive evolution can be modeled as an adaptive walk, in which each mutational step is taken with a fixation probability that has been derived by Kimura


⁹⁸


. We thus call the resulting adaptive walks Kimura walks (

Supplementary Methods 8

). We simulated 10

³

such adaptive walks starting from each of 15,000 randomly and uniformly selected starting (non-peak) TFBS genotypes from each landscape. Each random walk comprised up to 25 mutational fixation steps, unless it reached a fitness peak earlier. Each adaptive walk also accounted for known

E. coli

mutational biases


<sup>102

–

104</sup>


(

Supplementary Methods 8

).

As we hypothesized, most adaptive walks indeed reach only low peaks (90%, 85%, and 87% percent for the CRP, Fis, and IHF landscapes.). However, the percentage of walks that terminate at high peaks is several-fold greater than the proportion of high peaks. Specifically, 10 percent of walks terminate at high peaks in the CRP landscape, which is 3.6-fold higher than the percentage (2.8%) of high peaks in this landscape. In the Fis and IHF landscape, adaptive walks are 2 and 1.6-fold more likely to terminate at high peaks than expected from the proportion of these peaks among all peaks (7.4% and 8.1% percent) (

Figure 4a

).

To determine whether high regulatory peaks are more accessible due to chance alone, we compared the empirical landscapes to randomized “null” landscapes. Specifically, we generated these randomized landscapes by permuting regulation strengths across genotypes while preserving the sampled genotype network (

Supplementary Methods 7.6

). On each randomized landscape, we then performed adaptive-walks (15,000 walks for each of 10

³

starting genotypes) with the same parameters as for the biological landscape. For all three landscapes, the fraction of adaptive walks reaching high regulatory peaks in the empirical landscapes exceeds that for the randomized landscapes by almost three-fold, a difference that is statistically significant (

Supplementary Figure S30

,

Supplementary Methods 9

). In sum, rugged regulatory landscapes strongly constrain evolutionary trajectories, yet do not render the evolution of strong regulation vanishingly rare. Instead, strong regulatory phenotypes remain evolutionarily attainable at levels that exceed null expectations, even though they are reached by only a minority of evolving populations.

The adaptive walks that reached a high peak were short (

Figure 4b

). On average, they were also no more than half a mutational step longer than the shortest genetic distance between each starting genotype and peak (

Figure 4b

, CRP landscape: path length 3.2 ± 1.6 [mean±s.d.] vs. genetic distance 2.8 ± 1.2; Fis landscape: path length 3.1 ± 1.5 vs. genetic distance 2.7 ± 1.2; IHF landscape: path length 3.1 ± 1.5 vs. genetic distance 2.7 ± 1.2).

These short paths can be explained by our previous observation that peaks are widely distributed across the genotype spaces of the three landscapes. This distribution makes it easier for populations starting from any (non-peak) genotype to find a local peak through few mutations. Additionally, path lengths realized during adaptive walks tend to be shorter than the average lengths of accessible paths. (

Figure 4b

).

Because genetic drift can cause evolving populations to escape a low fitness peak and attain a higher fitness peak, we also asked how small population sizes affect the likelihood that a population attains a high fitness peak (

Supplementary Material 8

,

Supplementary Figures S26

-

28

). When simulating adaptive evolution in populations of N=10

²

individuals, we found that the likelihood that a population reaches a high peak increased by at least 10% percent (to 18%, 20%, and 21% for the CRP, Fis, and IHF landscapes,

Supplementary Figures S26

-

S28

).

We also studied how deviations from the strong selection weak mutation regime would affect evolutionary dynamics. In large populations or at high mutation rates, populations tend to be polymorphic and are subject to clonal interference, where the most advantageous of several co-occurring mutations typically dominates and becomes fixed


<sup>105

–

107</sup>


. To approximate this dynamic, we conducted simulations using “greedy” adaptive walks (

Supplementary Methods 8

)


<sup>25

,

108</sup>


. In such an adaptive walk, it is always a genotype’s mutational neighbor with the highest increase in regulation strength that becomes fixed


<sup>100

,

108

–

110</sup>


. In other words, a greedy walk starting from a given genotype is deterministic. We, therefore, simulated only one walk for each of the 15,000 randomly chosen (non-peak) starting genotypes. We found that the fraction of greedy walks reaching high regulatory peaks is slightly lower than in the SSWM regime, with 1%, 2%, and 5% fewer walks achieving such peaks in the CRP, Fis, and IHF landscapes, respectively (

Supplementary Figure S29

). This outcome is expected, because greedy walks prioritize immediate fitness gains at the expense of the ability to discover higher fitness peaks


¹⁰⁸


.

The attainment of peaks is highly contingent on chance events

Because different basins of attraction share many TFBS genotypes (

Figure 3d

,

Supplementary Figures S24

-

S25

), it is possible that adaptive evolution starting from any one genotype can reach different peaks, depending on which mutational paths it takes. In other words, the structure of the landscapes we study may lead to evolutionary contingency


<sup>111

,

112</sup>


– the dependence of an outcome of evolution on unpredictable chance events


<sup>47

,

111

,

113</sup>


. Indeed, non-peak genotypes can access on average around 30% of the total number of high peaks in each landscape (

Supplementary Figure S25

). To assess the prevalence of evolutionary contingency, we determined how many different peaks are attained by 10

³

adaptive walks starting from the same randomly chosen genotype. We performed this analysis on 15,000 starting genotypes, focusing on the subset of those starting genotypes from which at least one high peak is reached during the 10

³

walks. Specifically, 28.6%, 23%, and 24.5% of starting genotypes for the CRP, Fis, and IHF landscapes, reached at least one high peak. Notably, these percentages represent the fraction of starting genotypes that can access at least one high peak, not the total fraction of adaptive walks that lead to a high peak, which is lower (as shown in

Figure 4a

). For instance, while adaptive walks starting from 28.6% of genotypes reached at least one high peak in the CRP landscape only 10% of all adaptive walks in this landscape end at a high peak (

Figure 4a

). Importantly, for any one starting genotype where random walks reached at least one high peak, they typically reached more than one high peak (

Figure 4c

, median±IQR, 4±4, 4±8, 6±6 for the CRP, Fis, and IHF landscapes, respectively). The number of high peaks attained from any one starting genotype was as large as 21, 30, and 33 for the three respective landscapes. Thus, evolutionary contingency is pervasive in all three landscapes. In addition, not only the end point of evolution but also the evolutionary trajectories leading to this end point show contingency. That is, adaptive walks starting from a specific genotype and ending at a specific high peak often take multiple paths to this peak (

Supplementary Figure S31

).

Biases towards specific evolutionary outcomes have been documented in both empirical and computational studies


<sup>18

,

31

,

47

,

111

,

114

–

116</sup>


. They also exist in our landscapes (

Figure 4d

,

Supplementary Figure S31

).

Figure 4d

illustrates both contingency and bias for 10

³

adaptive walks starting from each of 10 different genotypes in the CRP landscape (thus a total of 10

⁴

adaptive walks). Depending on the starting variant, the adaptive walks reached between 1 and 16 high peaks. Whenever different peaks are reached, they are reached by markedly different proportions of adaptive walks. The same holds in the Fis and IHF landscapes (

Supplementary Figure S33

). Additionally, while multiple routes to each peak can be traversed (

Supplementary Figure S31

), we observed in all three landscapes biases towards specific paths that are taken more frequently than others (

Supplementary Figure S34

).

The number of these variants is shown as an integer on top of the panel, and its percentage among all 15,000 starting variants is shown in parentheses (color legend). Underneath it, the panel shows the mean ± 1 s.d. of the number of attained high peaks.

a. d. Some high peaks are reached more often than others.

We randomly and uniformly sampled 10 starting genotypes from the CRP landscape, started 10

³

adaptive walks from each, and recorded the number and frequency of distinct high peaks attained in these random walks. Results for each starting genotype are symbolized by a vertical bar. The number of stacks within each bar (delineated by horizontal lines, also indicated by an integer above each bar) indicates the number of high peaks reached by the 10

³

adaptive walks. Starting variants are ordered in ascending order based on this number of attained peaks. Stack height indicates the fraction of walks that reached the same peak, and is indicated in red, orange, and yellow for the three most frequently attained peaks.

See


Supplementary Figure S33

for the Fis and IHF landscapes.

Lastly, because sort-seq measurements are subject to experimental uncertainty


<sup>71

,

117

,

118</sup>


, an important question is whether such noise affects the inferred structure of a regulatory landscape and the resulting evolutionary dynamics. To address this issue we explicitly incorporated empirically estimated measurement uncertainty into our fitness comparisons and repeated pertinent analyses under two conditions: a noise-free landscape (𝒢


^S


) and an “uncertainty-aware” landscape 𝒢

^S,τ

using genotype-specific noise estimates (τ, see

Supplementary Figure S35

,

Supplementary Methods 10

). We examined whether noise-induced changes in landscape structure influence evolutionary trajectories. To do so, we compared adaptive walk dynamics between the two kinds of landscapes using identical population genetic parameters. Despite differences in peak counts and local topology, the overall pattern of genotype visitation during adaptive walks was highly similar between the two kinds landscapes. Specifically, the visitation frequency profiles of genotypes were strongly correlated between 𝒢


^S


and 𝒢


^S,τ


landscapes (Spearman’s ρ<0.001 shown in

Supplementary Figure S35

). This means that genotypes frequently accessed in the noise-free landscape remain frequently accessed in the noisy landscape.

Discussion

We evaluated the ability of three

E. coli

global regulators to control gene expression through each of more than 30,000 TFBSs for each regulator. To this end, we utilized a synthetic plasmid-based system that facilitates high-throughput fluorescence measurements. This system allowed us to quantify the regulation strength of individual TFBSs by measuring gene expression through GFP fluorescence


⁷⁴


. Additionally, the system insulates the control of transcription from any direct effects library sequences might have on transcription, such as the transcriptional and translational impacts of 5’ untranslated regions on gene expression


<sup>119

,

120</sup>


. Recent studies have investigated large empirical datasets of cis-regulatory genotypes, examining the impact of sequence variation on gene regulation in both eukaryotes and prokaryotes


<sup>6

,

16

,

72

,

81

,

121

–

124</sup>


. However, few studies have focused on these interactions from an evolutionary perspective and studied adaptive landscapes of gene regulation, as we do here


<sup>6

,

16

,

123</sup>


.

We showed that the regulatory landscapes of all three TFs are highly rugged and have multiple peaks. The ruggedness of all three landscapes is also supported by the prevalence of epistasis between pairs of TFBS mutations (

Supplementary Table S5

). A particularly important form of epistasis is sign epistasis


<sup>24

,

93

,

94</sup>


, because it can lead to multiple adaptive peaks


<sup>24

,

93

,

94</sup>


(see

Supplementary Methods 7.5

). Our landscapes contain up to 65% of mutation pairs with sign epistasis, a value that is especially high compared to the almost exclusively additive interactions of mutations in eukaryotic TFs


<sup>6

,

125</sup>


. However, the TFs we study are not exceptions among other prokaryotic TFs. Recent smaller-scale studies have shown that epistatic interactions are common in binding sites of local prokaryotic regulators, such as AraC (sign epistasis in more than 50% of 20 mutants


¹²⁶


) and Cl (sign epistasis in 85% of 113 mutants


¹²⁷


).

A possible reason for this greater incidence of epistasis lies in the nature of prokaryotic TFBSs. Specifically, prokaryotic TFBSs are at approximately 20bps twice as long as eukaryotic TFBSs


<sup>80

,

128</sup>


and exhibit symmetries that reflect the dimeric state of their cognate TFs


<sup>129

–

131</sup>


. These factors may increase the likelihood of intramolecular epistasis. Our observations raise important questions for future work, such as why the landscapes of prokaryotic TFBSs differ so dramatically from those of eukaryotic ones. And what do these differences imply for the evolutionary dynamics of gene regulation?

Despite the high ruggedness and pervasive epistasis of these landscapes, we found that a modest fraction of evolving populations can still access the highest regulatory peaks. Specifically, our evolutionary simulations show that 10% of populations with a size typical of

E. coli

reach one of the highest peaks. This percentage is significantly higher than in randomized landscapes (

Supplementary Methods 9

;

Supplementary Figure S30

), which shows that the structure of our regulatory landscapes facilitates access to stronger regulation than expected by chance. We speculate that this property reflects the biological role of global regulators, which coordinate the expression of many target genes


¹¹


and thus must operate across a wide spectrum of regulation strengths, while still permitting the evolution of strong regulation when necessary.

The clonal interference that may occur in even larger populations or in populations with high mutation rates reduces the accessibility of high peaks from 10% to 5% of evolving populations (

Supplementary Figure S29

). Conversely, in small populations where genetic drift can help a population escape from a low peak, this percentage increases to 18%. Numbers like these render the de novo evolution of strong transcription factor binding sites plausible for our three global regulators. Once such binding sites have originated in one population, they can also spread to others through horizontal gene transfer


<sup>132

,

133</sup>


.

In addition to increasing the ruggedness of a landscape, epistasis can also influence the evolution of populations on the landscape


<sup>24

,

94

,

134</sup>


. The regulation strength of a TFBS is partially determined by the combined interactions of the nucleotides within the TFBS


<sup>50

,

135

–

137</sup>


, which in turn affects TF-TFBS binding affinities. Different combinations of mutations can result in similar regulation strengths and create multiple evolutionary pathways to achieve optimal or near-optimal regulation


<sup>6

,

138</sup>


(

Supplementary Figure S31

). This can lead to overlapping basins of attraction among different peak TFBSs conveying strong regulation, such that even evolution starting from the same genotypes can take different paths and reach different peaks with similar regulation strengths. Indeed, we observed such overlapping basins in our landscapes (

Figure 3d

and

Supplementary Figure S24

). Moreover, epistasis can create plateaus of regulation strength where various combinations of nucleotides yield TFBSs with intermediate regulation strengths. These plateaus further contribute to the overlapping basins of attraction we observe, and may serve as common evolutionary intermediates that multiple starting sequences can traverse on their way to different peaks.

In a landscape with substantial epistasis, the sequence of mutations that occurs in an evolving population can also render adaptive evolution highly contingent on this sequence


<sup>111

,

112</sup>


. For example, some beneficial mutations within a TFBS might only be accessible after other specific mutations have occurred. Different starting sequences or early mutations can also change the spectrum of accessible mutations, leading to different peaks with similarly strong regulation


<sup>111

,

112</sup>


. Indeed, we observed in all three landscapes that different evolving populations starting from the same genotypes in the landscape attain different peaks (

Figures 3d

,

4d

and

Supplementary Figure S33

)


<sup>111

,

113

,

139

,

140</sup>


. Such contingency reduces the predictability of evolution


<sup>47

,

111

,

141</sup>


.

Despite the prevalence of contingency in all three landscapes, evolving populations starting from the same genotype more often attain some peaks than others (

Supplementary Figure S33

). Moreover, for each attained peak with multiple possible evolutionary routes, we also observed that some paths are more frequently transversed than their alternatives (

Supplementary Figure S34

). That is, evolution is biased towards traversing some paths and attaining some peaks more often than others


<sup>142

,

143</sup>


. These observations emphasize the complex interplay between chance, contingency, and evolutionary biases in shaping the outcomes of adaptive evolution.


<sup>47

,

111

,

113

,

115</sup>


.

The three TFs we studied here belong to different protein families, yet they have regulatory landscapes with similar topography. All three landscapes are highly rugged, highly epistatic, and harbor multiple peaks that are widely scattered, and this holds for peaks of low, intermediate, and high regulation strength. In addition, peak accessibility in all three landscapes increases with peak height (

Supplementary Figure S32

). On the one hand, these commonalities may be caused by common biological or biochemical properties of the three TFs. For example, CRP has been suggested to possess nucleoid-associated protein properties similar to Fis and IHF, due to its ability to bend and loop DNA


¹⁴⁴


. On the other hand, the commonalities may reflect general characteristics of global gene regulation. One of them is that global TFs often bind unspecifically to multiple TFBSs


<sup>11

,

13</sup>


. Also, they may bind DNA with a broad range of different affinities centered around intermediate affinity, rather than bind few sites but very strongly


¹³


. This possibility is supported by comprehensive analyses of in vitro eukaryotic TF binding affinities for thousands of TFBS variants, showing essentially continuous binding affinity distributions for hundreds of sites


⁶


.

Despite broad similarities among the three landscapes we study, we also found some differences. Most notable is the narrower distribution of regulation strengths for the binding sites of CRP compared to those of Fis and IHF (

Figure 2a

). This is not unexpected, given that CRP binding sites are known for being quasi-symmetric and less degenerate than those of Fis and IHF


<sup>57

,

60</sup>


. Previous studies have also shown that Fis and IHF binding sites are less conserved in their nucleotide sequence and more biased toward AT-rich content


<sup>43

,

145

,

146</sup>


. Differences in the distribution of regulation strengths may also result from differences in functions among the three proteins. CRP’s function is mostly restricted to gene regulation. As a result, it may have evolved to finely tune its regulatory output, resulting in a narrower distribution of regulation strengths. In contrast, Fis and IHF are involved in functions beyond gene regulation, such as DNA replication and genomic structural maintenance


<sup>147

,

148</sup>


, which might impose additional constraints or require a broader range of DNA binding strengths.

One limitation of our work stems from our rigorous quality filtering of sort-seq data. As a result, we lack regulatory data for some 40% of the TFBSs in each landscape. This limited diversity of reliable data is a common feature in mutational library studies. It has several technical causes, such as biases in library synthesis


¹⁴⁹


, PCR amplification


¹⁵⁰


, cloning, and loss of sequence diversity after cell sorting


¹²²


. In future work, this limitation could be overcome by a combination of strategies, such as subsampling complete genotype spaces or combining different molecular methods to overcome biases and diversity loss during PCR amplification


<sup>151

,

152</sup>


, sorting


<sup>71

,

117

,

153</sup>


, and high-throughput sequencing


¹²³


.

Importantly, although undersampling of genotype space is a limitation, standard approaches such as random subsampling or predictive modeling are not straightforward remedies. Several of our core analyses—including peak identification, quantification of epistasis, and assessment of evolutionary accessibility—rely on combinatorially complete local neighborhoods in genotype space. Random subsampling of genotypes would remove mutational neighbors, and thereby confound pairwise comparisons and the interpretation of landscape topology. Predictive modeling could be used to infer missing genotypes and reconstruct more complete landscapes


¹⁵⁴


, but it requires additional assumptions that introduce their own limitations. In addition, developing, validating, and benchmarking such models would be beyond the scope of this study, which is focused on empirical landscape mapping, and can serve as a starting point for future modeling work.

A second limitation comes from our use of the sort-seq method, which is best-suited for our work, because it allows the high-throughput measurement and sorting of millions of individual cells in a standardized and straightforward manner. However, the method’s accuracy depends on the binning procedure. Other studies have used between two and several dozen bins, with or without unbiased sampling


<sup>71

,

73

,

81

,

117

,

121

,

122

,

155</sup>


. Recommendations emerging from this work include sorting of cells into at least four logarithmically (log

₂

) equally-spaced bins, each covering approximately 12.5-15% of the fluorescence distribution,


<sup>81

,

122

,

156</sup>


. We followed these recommendations, using 13 bins. In addition, we computed a weighted average to calculate expression values from sequences appearing in multiple bins, a straightforward method validated by robust studies in transcriptional regulation analysis


<sup>16

,

72</sup>


. In addition, we validated individual regulation strengths with an independent method to demonstrate its reliability.

A third limitation of our study is that we only examined variation at eight positions for each TF. We selected these positions based on their importance for DNA binding and regulation by our TFs, as evidenced by their high information content. We cannot exclude the possibility that selecting a different set of positions could yield different landscape topographies. However, we speculate that less information-rich position would not reduce but rather increase the potential for the de novo evolution of strong TFBSs. For example, they may facilitate landscape navigability through extradimensional bypasses


<sup>96

,

157</sup>


or provide small-incremental changes in regulation strengths. These small changes help populations reach peaks via diminishing returns effects, meaning that as a population gets closer to a peak, each subsequent mutation contributes progressively smaller improvements to regulation strength


<sup>158

,

159</sup>


. Investigating a wider array of positions and larger regulatory landscapes remains an important task for future work.

Fourth, we use a simplified empirical system for the study of gene regulation—a promoter followed by a single TFBS. Although this regulatory architecture exists for some genes


¹⁶⁰


, global regulators often form part of more complex, combinatorial architectures


<sup>4

,

160

,

161</sup>


that are influenced by environmental factors


<sup>2

,

162

,

163</sup>


, concentrations of active TFs


<sup>164

–

166</sup>


, chromosome structure


<sup>167

–

171</sup>


, methylation states


¹⁷²


, and more. Studies on more complex promoter architectures may reveal regulatory landscapes with different topographies.

Lastly, we also analyzed the adaptive evolution of TFBSs in a simplified manner, performing data-driven evolutionary simulations rather than experimental evolution. Although many studies assume that regulation strength and fitness are correlated, this is not always the case. Low binding affinities can be adaptive during development


¹⁷³


, and many genes exhibit a nonlinear fitness-expression function


¹⁷⁴


with a plateau of maximal fitness across a wide range of expression levels


<sup>175

–

177</sup>


. For instance, most mutations and polymorphisms in the promoter of the yeast gene TDH3 do not significantly affect fitness in a glucose-rich medium


¹⁷⁶


. For these reasons we focused our observations and interpretations on regulatory phenotypes (regulation strength). Our simplified approach allowed us to model population evolution in a large genotype space and avoid monitoring thousands of evolving regulatory sequences simultaneously in vivo


<sup>23

,

178</sup>


. However, it cannot replace experimental evolution of TFBSs, which also remains an important challenge for future work.

Transcription factor binding sites are among the simplest units of biological organization. Our work provides the first large-scale analysis of the regulatory landscapes formed by such sites for global transcriptional regulators. It shows that strong binding sites can readily evolve de novo, even though prokaryotic transcription factor binding sites are much larger than their eukaryotic counterparts and have more rugged regulatory landscapes. In addition, the evolution of these simple sequences also displays phenomena that have been characterized in much more complex systems, such as evolutionary contingency and evolutionary biases.

Materials and methods

The Supplementary Materials contain extended details of experimental procedures and data analysis.

Strains and plasmids

Bacterial strains and plasmids used in this work are listed in

Table S1

. We obtained electrocompetent

E. coli

cells of strain SIG10-MAX

^®

from Sigma Aldrich (CMC0004). We used this strain for molecular cloning and library generation due to its high transformation efficiency. The genotype of this strain (

Supplementary Table S4

) is similar to DH5α (Sigma Aldrich commercial information, see

Supplementary Table S4

). The strain is resistant to the antibiotic streptomycin.

We amplified plasmid libraries in SIG10-MAX

^®

, extracted their DNA, and transformed them into mutants derived from

E. coli

K-12 strain BW25113 that harbor chromosomal deletions of the

crp

,

fis

, or

ihfa

gene. We obtained these mutant strains from the KEIO collection


¹⁷⁹


and used them for sort-seq experiments. The design, genetic parts, and assembly of the plasmid vectors we used in this study are available in the Supplementary material, as are all primers, TFBS sequences/libraries, strains, and plasmids.

Sort-Seq procedure

To explore the regulatory effects of each transcription factor on binding sites in the corresponding library, we constructed three plasmids, each of which enables the inducible expression of one of our three TFs. These are plasmids pCAW-Sort-Seq-V2-CRP, pCAW-Sort-Seq-V2-Fis, and pCAW-Sort-Seq-V2-IHF (

Supplementary Methods 3

and

Supplementary Table S1

). We cloned the TFBS libraries (

Supplementary Table S3

) into their respective plasmids and then transformed them into mutant strains lacking the corresponding TFs (

Δcrp

,

Δfis

, and

Δihf

, as listed in

Supplementary Table S2

). We induced TF expression using anhydrotetracycline (Atc) and, after overnight growth, performed cell sorting for cell populations. During sorting, we distributed cells into 13 equally-spaced logarithmic bins based on their fluorescence levels. We replicated each sort-seq experiment three times from three separate library transformations for each TF. To mitigate the impact of extrinsic noise (gene expression variation among cells


¹⁸⁰


), we adhered to standard protocols by normalizing our GFP fluorescence measurements against mScarlet-I fluorescence values obtained from flow-cytometry assays


<sup>7

,

71

,

181

,

182</sup>


. We subsequently recovered the sorted cells from each bin in 50mL Falcon tubes containing 10 mL of LB medium supplemented with chloramphenicol and incubated them overnight at 37°C with shaking at 220 rpm. After this growth period, we re-sorted cell cultures from each bin to eliminate potential contaminants and ensure that the cell populations had preserved their fluorescence distributions. Following re-sorting, we extracted plasmids from the cell population of each bin. We amplified and barcoded the TFBS region from each population through a polymerase chain reaction (PCR). Lastly, we sequenced barcoded amplicons containing TFBS sequences, and used the sequencing results to calculate the regulation strengths of each TF to its TFBSs in the corresponding library. More details are provided in the Supplementary Material.

Regulation strengths

Due to gene expression and measurement noise, individual TFBS variants in a sort-seq experiment usually appear in more than a single bin, and their read count (frequency) varies among bins


<sup>71

,

72

,

121

,

153</sup>


. Following established practice


<sup>18

,

72

,

123</sup>


, we used a weighted average of these frequencies for each variant to represent the mean expression level caused by the variant. To facilitate the interpretation of this quantity, we converted this expression level into a regulation strength relative to the highest observed regulation strength for a given TF, to which we assigned a value of one. From each library, we selected a single naturally occurring binding site for each TF that was previously characterized in the literature as a strong binder. We called this TFBS the WT sequence and used it as a baseline to separate weakly (higher GFP expression) from strongly (lower GFP expression) regulating TFBSs.

Validating regulation strengths with plate reader measurements

To further validate our regulation strength data, we chose 10 DNA binding sites from each bin (10 variants ×13 bins = 130 variants in total per TF library,

Supplementary File 2

), covering a wide range of measured regulation strengths. We cloned these sequences into the appropriate vector pCAW-Sort-Seq-V2-TF (TF: CRP, Fis or IHF), and transformed them into the appropriate mutant strain. We picked individual colonies and grew them overnight (16 hours, 37°C, 220 rpm) in liquid LB supplemented with 50 μg/mL of chloramphenicol and anhydrotetracycline. We diluted the cultures to 1:10 (v/v) in cold Dulbecco’s PBS (Sigma-Aldrich #D8537) to a final volume of 1 mL. We transferred 200 μl of the diluted cultures into individual wells in 96-well plates and measured GFP fluorescence (emission: 485nm/excitation: 510nm, bandpass: 20nm, gain: 50), as well as the optical density at 600nm (OD

₆₀₀

) as an indicator of cell density. We then normalized fluorescence by the measured OD

₆₀₀

value to account for differences in cell density among cultures and compared the obtained ratios to the previously inferred regulation strengths for the 130 selected variants. We performed all such measurements in biological and technical triplicates (three colonies per sample, and three wells per colony, respectively).

Supplementary methods

1.
General procedures

Although all general procedures have been previously described


¹⁸


, we describe them again below for completeness.

1.1
Media and reagents

To prepare SOB medium, we mixed 25.5g of solid medium stock (VWR J906) with 960 ml of purified water and subjected the resulting suspension to autoclaving. To prepare the SOC medium, we dissolved 20 ml of 1 M D-glucose (Sigma G8270) and 20 ml of 1 M magnesium sulfate (Sigma 230391) in 960 ml of pre-prepared SOB solution. For the LB medium, we combined 25g of solid medium stock (Sigma-Aldrich L3522) with 1 liter of purified water and then autoclaved it. To prepare the M9 minimal medium, we diluted M9 minimal salt sourced from Sigma (M6030) in distilled water as per the manufacturer’s guidelines, autoclaved the solution, and added 0.4% glucose (Sigma G8270), 0.2% casamino acid (Merk Millipore, 2240), 2 mM magnesium sulfate (Sigma 230391), and 0.1 mM calcium chloride (Sigma C7902). Where required, we supplemented growth media with chloramphenicol (50 µg/mL), anhydrotetracycline (100 ng/mL, Cayman-chemicals #10009542), and/or glucose (0.4% w/v final concentration). We prepared anhydrotetracycline by diluting the dried chemical in absolute ethanol, from a stock concentration (1000X) of 100 µg/mL to a working concentration of 100 ng/mL.

1.2
Overnight incubation of cultures in liquid and solid medium

We cultivated bacteria in liquid LB medium (using either 15mL or 50mL Falcon tubes), enriched with chloramphenicol at a concentration of 50 µg/mL. We incubated these cultures for a period of 16 hours at a temperature of 37°C, with a shaking speed of 200rpm and 50 mm orbital motion, using an Infors HT Multitron Incubator Shaker. Similarly, for cultures in solid medium, we grew bacterial colonies on LB-agar plates (using sterile plastic Petri dishes of 90mm × 15mm dimensions), also supplemented with chloramphenicol at a concentration of 50 µg/mL, and incubated them for the same time and at the same temperature.

1.3
PCR Reactions

Except where specifically mentioned, we amplified DNA fragments through a polymerase chain reactions (PCR), employing Q5

^®

high-fidelity polymerase (NEB #M0491L) to minimize mutation introduction into the amplicons. We followed the protocol recommended by NEB, aiming for a final reaction mixture of 50uL. We conducted each PCR reaction twice, and combined the products from these duplicates after the completion of the reaction. We determined the primer melting temperatures (Tm) using the NEB Tm calculator (accessible at

), with primers at a concentration of 500nM.

1.4
Verifying PCR products through gel electrophoresis

Unless indicated otherwise, we verified successful PCR amplification via gel electrophoresis to ensure the presence of singular-band amplicons and the absence of non-specific bands. This process involved separating PCR products in a 0.8% agarose Tris-EDTA (TAE) gel. We conducted electrophoresis for 45 minutes at 120V, or until the bands had progressed beyond the halfway point of the gel’s total length.

1.5
DNA purification with commercial kits

Once we had confirmed a successful PCR through gel electrophoresis, we purified the PCR products utilizing the Monarch

^®

DNA PCR/Gel Extraction Kit (NEB #T1020L), adhering to the manufacturer’s protocol. In situations requiring further purification (such as the occurrence of non-specific bands post-PCR), we performed gel purification. This involved mixing 10 μL of 6x NEB DNA dye with each 50 μL PCR product, then loading all 60 μL onto a 1% agarose gel. We carried out electrophoresis for 45 minutes at 120V, or until the bands had moved more than halfway through the gel. We then excised the DNA band corresponding to the amplified sequence using a scalpel. For extracting DNA from the gel, we used the Monarch

^®

DNA Gel Extraction Kit (NEB #T1020L).

1.6
Gibson assembly


¹⁸³

We assembled PCR-amplified fragments using the NEBuilder-HiFi

^®

DNA Assembly Master Mix kit (NEB #E2621L). We determined the molarity required for assembly based on the guidelines outlined by the Barrick Lab (details available at

). We incubated the assembly mix for one hour at 50°C in a dry bath incubator, followed by cooling on ice for subsequent steps.

1.7
Preparation of electrocompetent cells

We utilized glycerol/mannitol step centrifugation to prepare electrocompetent cells


¹⁸⁴


. To this end, we first cultured the appropriate

E. coli

strain (

Supplementary Table S4

) in 5 mL SOB medium at 37°C with a shaking speed of 250 rpm overnight. The next day, we transferred 3 mL of the culture to 300 mL SOB medium, and incubated under the same conditions until the OD600 reached a value between 0.4 and 0.6 (measured at an optical path length of 1 cm), which took approximately 2-4 hours. After cooling the culture on ice for 15 minutes, we centrifuged the cells at 4°C and 1,500 g for 15 minutes. We then resuspended the cells in 60 mL of ice-cold distilled water (dH

₂

O) and divided them into three 50 mL tubes. Gradually, we added 10 mL of an ice-cold glycerol/mannitol solution (consisting of 20% glycerol (w/v) and 1.5% mannitol (w/v)) to each tube using a 10 mL pipette. We centrifuged the tubes at 1,500 g and 4°C for 15 minutes in an Eppendorf 5810/5810 R centrifuge with acceleration/deceleration set to zero. After discarding the supernatant, we resuspended cells in 3.0 mL of the same glycerol/mannitol solution. We transferred these suspensions to pre-cooled 1.5 mL tubes, and incubated them in a dry ice-ethanol bath for about 1 minute. Finally, we stored the suspensions at -80°C for future transformation experiments.

1.8
Electroporation

In all transformation experiments described in this study, we used 100 µL of electrocompetent cells for electroporation, employing 0.2 cm cuvettes (EP202, Cell Projects, UK) and a Micropulser electroporator (Bio-Rad) set to the EC3 setting (15k V/cm). Post-electroporation, we recovered cells in 1 mL of SOC media, warmed in advance in 15 mL Falcon tubes. This recovery step lasted for 1.5 hours at 37°C with a shaking speed of 220 rpm. Unless specified otherwise, we spread 300 µL of each recovered culture on an LB agar plate that contained 50 μg/mL chloramphenicol. We then incubated this plate overnight for 16 hours at 37 °C. Subsequently, we confirmed the identity of the clones on the plate via Sanger sequencing.

2.
The design of plasmid pCAW-Sort-Seq-V2

The plasmid pCAW-Sort-Seq-V2 is a derivative of the plasmid pCAW-Sort-Seq


¹⁸


. Briefly, plasmid pCAW-Sort-Seq harbours a pBBR1 replication origin, which ensures a broad host range and a low copy number—typically between 5 to 10 copies per cell


¹⁸⁵


.It also harbours a chloramphenicol resistance gene, a TetR repression system


¹⁸⁶


and a TFBS measuring module. The latter consists of an interchangeable TFBS that lies between a constitutive promoter and a superfolder GFP (

sfgfp



¹⁸⁷


) reporter gene. In this system, TF-TFBS interactions can decrease GFP production by physically obstructing the bacterial RNA polymerase, a phenomenon known as steric hindrance. The reporter gene

sfgfp

is insulated by a transcriptional insulator named RiboJ, a synthetic ribozyme that removes 5’UTR interferences from variable TFBS sequences in the mRNA by self-cleavage


¹⁸⁸


. More details and features have been previously described in ref.


¹⁸


. Here, we have integrated a bicistronic expression cassette into pCAW-Sort-Seq to enable the regulated expression of a TF of choice. This cassette harbors the gene encoding one of our focal TFs situated upstream of a

mscarlet-I



¹⁸⁹


reporter gene to monitor the expression of this bicistronic operon via fluorescence. Regulation is achieved through the

pLtetO-1



¹⁸⁶


promoter, a synthetic promoter tightly repressed by TetR. Expression is initiated by the addition of anhydrotetracycline (Cayman Chemicals, catalog #10009542), which relieves TetR-mediated repression and activates the expression of the entire cassette.

3.
Construction of the plasmid pCAW-Sort-Seq-V2 and its variants

We initiated the construction of pCAW-Sort-Seq-V2 by synthesizing (Twist Biosciences, Calfornia, USA) and cloning a codon-optimized

mscarlet-I

gene expression cassette into the pCAW-Sort-Seq plasmid. This gene is oriented antiparallel to the tetracycline resistance gene (

tetr

). This cloning step resulted in the formation of the intermediate pCAW-Sort-Seq-V2t plasmid (

Supplementary Figure S1

). Subsequently, we amplified for each of our three TFs the encoding gene (

crp

,

fis

and

ihf

) from the genome of the

E. coli

strain BW25113 and inserted it upstream of the

mscarlet-I

gene in a bicistronic operon configuration using Gibson assembly. This process produced TF-specific expression plasmid variants (

Supplementary Figure S2

). The final phase involved removing the core promoter to create negative controls for each TF and cloning libraries into each plasmid variant upstream of the sf

gfp

gene. This yielded the plasmids used in our experiments (

Supplementary Table S1

,

Supplementary Figure S2

). The specific sets of primers used for each PCR reaction are listed in

Supplementary Table S6

.

4.
Library design, synthesis, and cloning

We based the design of the TFBS library for each TF on consensus sequences available in the RegulonDB database


⁷⁰


, the most comprehensive database for

E. coli

transcriptional studies. We designed each library by randomizing the eight most important TFBS positions, such that each of the four nucleotides had an equal probability (0.25) to occur at each position. The expected library size for this approach is (4

⁸

)

⁼

65,536 sequences. Library compositions can be found in

Supplementary Table S3

. We designed libraries in the Snapgene

^®

software (

), and had them synthesized by IDT (Coralville, USA) as single-stranded DNA Ultramers

^®

of 140bp (4nmol). We resuspended each library in nuclease-free distilled water and serially diluted it to a concentration of 50ng/uL. We used Ultramers

^®

as templates in a PCR reaction for the formation of dsDNA fragments and library amplification. We amplified the Ultramer

^®

DNA molecules with the following PCR program: 98°C/30 s; 25 cycles of 98°C/10 s, 60°C/15 s and 72°C/80 s; and 1 cycle of 72°C/5 min. We opted for a maximum of 20 cycles to reduce amplification biases. We analyzed amplification products through gel electrophoresis to confirm that only a single product band with the expected size of around 140bp was present for each PCR reaction. After confirming the presence of single bands, we purified the products using the Monarch

^®

DNA gel extraction kit (NEB #T1020L). Whenever unspecific bands appeared during electrophoresis, we gel-purified the PCR products using the same kit.

For the construction of each plasmid-based library, we digested 1μg of the purified library with HindIII-HF (NEB #R3104) and BamHI (NEB #R3136) restriction enzymes in a 100μL reaction, followed by overnight incubation at 37°C. We isolated the appropriate cloning plasmid from its host strain using the QIAprep spin miniprep kit (Qiagen, Germany), and digested it with the same enzymes. Post-digestion, we added 3μL of Quick CIP (calf intestinal alkaline phosphatase) to prevent self-ligation by dephosphorylating DNA ends. We purified the ligated DNA using the Monarch

^®

DNA gel extraction kit (NEB #T1020L).

We performed the ligation with a 10:1 molar ratio of insert-to-vector, using 100ng of vector, 10 units of T4 DNA ligase (NEB #M0202L), and 2 µL of 10X ligation buffer in a 20 µL reaction. We incubated the mixture at 20-22°C for approximately 16 hours, followed by 15-minute inactivation of the ligase at 65°C. We purified the ligation product, resulting in 10 µL of DNA resuspended in dH

₂

O. We transformed

E. coli

SIG10-MAX® cells by electroporation with this purified product.

Post-transformation, we plated 50 µL of each recovered culture on LB agar for colony-forming unit counting (cfu) and transformation efficiency estimation. From the agar plates, we selected 30 colonies for colony PCR and Sanger sequencing to assess library diversity (NightSeq

^®

service, Microsynth, Switzerland). Our mean transformation efficiency was 10

⁶

cells per transformation. We diluted the remaining 950 µL of transformants in 9 mL of LB medium with chloramphenicol and cultured it overnight. We then aliquoted the overnight culture into 1 mL cryotubes with 20% glycerol and stored at -80°C.

After transforming plasmid libraries into the SIG10-MAX® strains for reasons of transformation efficiency and plasmid maintenance, we extracted the plasmids using a QIAprep spin miniprep kit (Qiagen, Germany), and transformed them into the appropriate host mutant host strains


Δ



crp,

Δ

fis and

Δ

ihf

(see the strain genotypes in

Supplementary Table S2

). We cultured each library-transformed strain overnight, aliquoted it in 1 mL cryotubes with 20% glycerol, and stored it at -80°C for further experimentation.

5.
Analysing and sorting cells

In preparation for cell sorting, we cultivated cells harboring each library in liquid LB medium enriched with chloramphenicol. Specifically, we cultured 1mL of transformed cell aliquots and a streak of cells with a control plasmid (a promoterless pCAW-Sort-Seq-V2 plasmid lacking sfGFP expression) in 50ml Falcon tubes with 9mL LB medium (containing 50 µg/mL chloramphenicol) overnight. Subsequently, we diluted these overnight cultures at a 1:100 ratio (v/v) in LB medium with chloramphenicol and divided them into two aliquots. We supplemented one aliquot with the anhydrotetracycline (Atc) inducer to express the plasmid-encoded TF. The other aliquot remained unchanged. We incubated both cultures for 5 hours until late-exponential/early-stationary phase (200RPM, 37°C). Subsequently, we diluted 20µL of the cultures in 1mL of cold filtered Dulbecco’s PBS (Sigma-Aldrich #D8537) in 15 mL FACS tubes.

We performed FACS-sorting on a FACS Aria III flow cytometer (BD Biosciences, San Jose, CA) using a 70 µm nozzle. We utilized a 488 nm laser for detecting forward scatter (FSC) and side scatter (SSC) with a 488nm/10nm band-pass filter. We set the flow rate to 1.0, adjusting sample dilution as necessary to achieve no more than ≈10,000 events/second. Considering the small size of bacterial cells, we reduced the particle detection threshold to the lowest feasible setting (200 arbitrary units on FSC and SSC channels), increasing it to a maximum of 500 units if background noise was excessive. We then adjusted FSC-H and SSC-H for cells with the negative control plasmid to center the bacterial population in the cytometer’s software (BD FACSDiva™ Software v9.0). We based the sorting and binning of cells on both mScarlet-I (PE-Texas-Red channel, excitation laser: YellowGreen 561nm, LP filter: 600 nm, BP filter: 610/20nm) and sfGFP fluorescence (FITC channel, excitation laser: 488nm, LP filter: 502nm, BP filter: 530nm/30nm), setting both the PE-Texas-Red and FITC channels voltages so that the median fluorescence of the negative control was between 0 and 100 (arbitrary units) on the FITC-H and PE-Texas-Red-H axes.

Initially, the sorting process involved establishing a gate for identifying cells that are red-fluorescence-positive, i.e., reporter and thus TF-expressing. To establish this gate, we began by measuring the autofluorescence of our negative control culture on the PE-Texas-Red-H axis. Thereafter, we examined the fluorescence of a positive control that had been induced with Atc and was expressing the mScarlet-I protein. We then configured a gate around the positive mScarlet-I-expressing population, ensuring that all cells sorted through this gate exhibited a level of reporter expression higher than the negative control.

Next, we established the green-fluorescent sorting gates. For setting sorting gates on the FITC-H axis, we first recorded the autofluorescence of the negative control culture. This median autofluorescence defined the upper boundary of the lowest bin (B1) for the experimental population. We then analyzed the fluorescence of 10

⁶

cells expressing sfGFP and containing the library, without sorting, in order to establish the boundaries of the next binning gates. We took the lower bound of the highest bin (B13) for the experimental population to correspond to the 95th percentile of the fluorescence distribution of this population. We chose boundaries between the remaining 11 intermediate bins with equidistant spacing on a binary logarithmic (log

₂

) scale. After we had determined the gates in this way, we calculated the fractions of the previously 10

⁶

recorded cells that fell inside each bin.

To maintain statistical robustness and minimize sampling error in our downstream analyses, we wanted to ensure that each of the 65,536 unique sequences in our library was represented by at least 30 cells after sorting. In addition, it was crucial to account for an anticipated diversity loss of up to 70% of the cell population during sorting, due to factors such as cell death and the dilution of low-frequency and lower-fitness genotypes during the post-sorting recovery phase


¹²²


. To establish the number of cells required to be sorted initially, we thus determined a multiplicative factor 𝑓 based on the minimally needed number of 30 cells per sequence and the expected cell retention rate post-sorting of 30%, i.e.,

cells/sequence. We thus multiplied our initial library size by 100 for sorting purposes, i.e., we sorted total of 6,553,600 cells. This calculation aims to ensure that despite a substantial reduction in cell numbers post-sorting, each unique sequence remains adequately represented in the surviving cell population. We sorted cells into 1.5 mL Eppendorf tubes, each containing 500 µL of LB medium, and kept at 4 °C to prevent growth during sorting and sample processing. We replicated the entire sorting procedure three times, each based on independent library transformations.

After sorting, we added 1mL of LB without antibiotics to each tube and removed 20 uL of the resulting volume for serial dilutions. We transferred the remaining liquid culture (980 uL) to 50 mL falcon tubes and allowed each culture to recover for 2 hours (37°C, 220 rpm). After recovery, we added 9mL of LB supplemented with chloramphenicol, and grew the cultures overnight for freezing part of them in glycerol stock aliquots, for validating our binning procedure (see below), and for extracting plasmid DNA for subsequent PCR and sequencing steps. We used the 20 uL initially removed from each 1mL culture for preparing two serial dilutions (10

^-4

and 10

^-6

) that we plated on LB-Cm agar plates (200uL per plate) to estimate the post-sorting viability through colony forming unit (cfu) counting. By knowing how many cells were sorted into each bin, we can estimate how many cells would be expected in our dilutions and compare this number with the cfu counts we observed. In this way, we estimated that on average (across bins) 77% of cells remained viable (standard deviation: 18.15%). We also estimated the genetic diversity of the library through Sanger sequencing of DNA from individual colonies (NightSeq

^®

service, Microsynth, Switzerland).

To validate our binning procedure, we re-grew binned cultures from either overnight recovered cultures or frozen aliquot stocks. We then measured their expression distributions by flow cytometry, which reproduced the original expression measurements. This approach allowed us to compare the fluorescence distributions of the re-grown cultures with the pre-sorting distributions. Specifically, we assessed whether the geometric mean of the fluorescence distribution for each sorting bin matched those recorded during the initial sorting procedure. Next, we re-sorted cells derived from each bin in order to eliminate cell cross-contaminations and other factors that could alter the bin distributions. The geometric mean is often preferred over the arithmetic mean in this type of analysis, because it is less affected by the presence of outliers in the data


<sup>190

,

191</sup>


. It is also a more accurate representation of the central tendency of data that is log-normally distributed, which is often the case with flow cytometry fluorescence measurements


<sup>190

,

191</sup>


.

6.
DNA extraction and sequencing

We diluted 500 uL of individual glycerol stocks of each replicate subpopulation of sorted cells (i.e., cells from each bin of fluorescence intensity) in 5mL of LB supplemented with chloramphenicol in 15mL Falcon tubes, and grew the resulting cell culture overnight (16 hours, 37°C, 220 rpm). On the next day, we isolated plasmids from each culture using a QIAprep

^®

spin miniprep kit (Qiagen, Germany). In order to allow the sequencing of multiple pooled samples (multiplexing), we barcoded our regulatory region through PCR with specific HPLC-purified primers (

Supplementary Table S6

) provided by Eurofins (Konstanz, Germany). We added barcodes to the 5’region of the amplicon through a PCR reaction. We performed this PCR reaction with the Q5 high-fidelity polymerase, and did so in triplicate for each miniprep-isolated plasmid library. To calculate primer melting temperatures (Tm), we used the NEB Tm calculator (

) for a primer concentration of 500nM. We performed the PCR with the following program: 98°C/30 s; 25 cycles of 98°C/10 s, 64°C/30 s and 72°C/30 s; and 1 cycle of 72°C/2 min.

After PCR amplification, we digested the reaction products with the restriction enzymes DpnI (NEB #R0176L) and Exonuclease I (NEB #M0293L) in order to remove traces of genomic DNA, plasmids, and single-stranded DNA that could interfere with sequencing. The Master Mix we used for a single digestion harbored 1µL of 10x CutSmart

^®

Buffer (NEB #B6004S), 1µL of Exonuclease I (NEB #M0293L), 1µL of DpnI restriction enzyme (NEB #R0176L), and 7µL of distilled nuclease-free water. For each PCR product, we added 10µL of the Master Mix. We incubated the reaction for 1 hour at 37°C, following 15 minutes at 80°C for deactivation of the enzymes.

We purified the digestion products using the Monarch

^®

DNA PCR/Gel Extraction Kit (NEB #T1020L) and fractionated them through gel electrophoresis to confirm that only a single band with a size ≈150bp was present. After this confirmation, we pooled the purified PCR products from the different bins of each replicate sorting equimolarly to a total mass of 2,600 ng and a volume of 100 µL (26ng/µL of DNA) in 1.5mL Eppendorf tubes. We then sent the pooled purified amplicons for adapter ligation and sequencing at Eurofins (NGSelect Amplicons

^®

on Illumina HiSeq), obtaining 15 million paired-end reads (2 × 150 bp) for all samples.

We quantified the total number of reads (

t

) required for sequencing based on several factors, including the number of bins (

b)

, biological replicates (

r)

, strains (

s)

, genotypes (

g)

, and the reads per genotype (

p)

. Using these variables, we first calculated the total number 𝑛

_’(

of needed barcodes as follows:

For each of our three TFs, the values of these parameters are

b

=13 bins,

r

=3 biological replicates, and

s

=1 strains, yielding

n

_bc


=39. The total number of required sequence reads can be estimated through the equation:

where we aimed for

p

=30 paired-reads per genotype. The number

g

of expected genotypes per library is the same (65,536) for each TF. These values lead to a total of

t

= 3 × 65,536 × 30 = 7,667,712 required paired-end reads for each library.

7.
Data analysis

7.1
Filtering and preparing sequencing reads

We processed the sequencing data with a blend of custom python and awk scripts, complemented by established bioinformatics utilities. Firstly, we trimmed sequences by computationally excising Illumina adapters, followed by merging paired-end reads. We then organized the paired-end reads into distinct files, each tagged with a sequence barcode identifying the sequence bin from which the corresponding reads originated.

For the removal of Illumina adapter sequences from the paired-end reads, we employed Cutadapt


¹⁹²


with the following parameters:

Here, “ADAPTER_FWD” and “ADAPTER_REV_RC” are placeholders for the actual adapter sequences used, which are 5’-
AGATCGGAAGAGCACACGTCTGAACTCCAGTCA
-3’ for read 1, and 5’-
AGATCGGAAGAGCGTCGTGTAGGGAAAGAGTGT
-3’ for read 2.

Because our amplicons were short (153 bp) and each sequencing sample was divided into two files with overlapping single-end reads, merging the two files (single-end reads) was necessary, for which we used the FLASH software


¹⁹³


. After the filtering and merging of reads, we demultiplexed paired-end reads with the following FLASH parameters:

Next, we utilized the FastX toolkit (

)


¹⁹⁴


to discern and preserve only those sequences that exceeded a high-quality threshold of Q = 33. Subsequently, we refined our dataset by removing sequences that contained undesirable mutations or insertions/deletions (indels) within the region extending from the start of the core promoter to the end of the variable TFBS library. We performed this filtering step using a custom R


¹⁹⁵


script. Thereafter, we used custom awk and R


¹⁹⁵


scripts to convert the data into a table. Each row of this table contains data from one TFBS in the library, and each column contains the number of reads for this TFBS from one of the thirteen bins into which we had sorted cells. We estimated the average regulation strength of each TFBS from this data.

7.2
Calculating regulation strengths

We calculated regulation strengths as previously described


¹⁸


. Briefly, in sort-seq experiments, sequences often appear in multiple fluorescence bins due to random mis-sorting


<sup>71

,

117</sup>


and the stochastic nature of gene expression


¹⁸⁰


. Following methods established in previous research


<sup>16

,

72

,

123</sup>


, we determined the reporter expression level driven from each TFBS variant by calculating a weighted average of bins in which the variant occurred. This involved multiplying the frequency of each sequence (

x

_i


) in a given bin

i

by a numerical value representing that bin (

w

_i


=1,2,3,…,13), and then averaging these products over the total sequence count. Mathematically, this weighted average calculates as

where

e

is the expression level driven by the sequence.

This approach yields a continuous spectrum of expression levels

e

within the range of 1 to 13. High expression values indicate robust GFP expression and, thus, weak binding of a TF to a TFBS variant. To facilitate interpretation, we inverted this scale so that higher values indicate stronger repression and lower GFP expression, i.e., we define the regulation strength

b

(strength of regulation) as follows:

where

e

_max


=13 is the maximal expression level (maximal bin value). We then normalized

b

by the maximal regulation strength among all TFBSs we studied for a given TF. The resulting normalized regulation strength scores

b

vary from 0 to 1, where low scores denote TFBS variants with weak TF binding (weak reporter repression), and high scores indicate variants with strong TF binding (strong reporter repression). A score of

b

=1 signifies the highest regulation strength (repression) calculated among all TFBSs for a given TF.

7.3
Combining data from triplicates

As a quality-filtering step, we eliminated TFBS variants that did not appear in all three replicates or that were represented by less than 30 reads in total (summing their read counts over the thirteen bins). Given that we have 13 bins and most sequences are typically found within an average of 3.4 bins, we wanted a minimum of 10 reads per bin to prevent misinterpretation of repression levels and also guide our threshold’s choice. This procedure reduced the fraction of TFBS variants for which we had sort-seq-based sequence data from 95%, 90%, and 93% of the total library size (65,536 sequences) for CRP, Fis and IHF, respectively, to 49%, 66% and 63%.

Subsequently, we calculated the regulation strengths

b

of the remaining TFBS variants (

Supplementary Materials and Methods 7.2

). Then, we determined the coefficient of variation of regulation strengths across replicates for each TFBS variant, which estimate the consistency of measured regulation strengths among replicates. Notably, most TFBS variants exhibited a coefficient of variation (CV) below 0.5, which was the threshold we set for filtering sequences. Sequences with a CV above 0.5 were excluded from the analysis. This threshold is commonly used in transcriptional studies employing fluorescent reporters to account for transcriptional noise and measurement variability


¹⁸⁰


. Finally, we averaged regulation strengths for each TFBS variant across all replicates and normalized the resulting averages by the maximal observed regulation strength among all TFBSs of a given TF.

7.4
Frequency matrices and sequence logos

We generated frequency matrices of nucleotides that occur in TFBSs from each fluorescence bin by counting the frequency of each nucleotide at each variable position of the TFBS library. From this data, we generated heatmaps and DNA sequence logos representing the frequency matrices graphically. A sequence logo consists of a stack of the letters A, C, G, and T, at each position of a DNA sequence, where the relative size of each letter indicates its frequency in the sequence. The total height of the stack corresponds to the information content of that position, in bits


<sup>49

,

196</sup>


. A sequence logo is a graphical representation of the informational properties of a TFBS. When mutated, nucleotides with high information content are more likely to lead to a loss of binding (repression) than nucleotides with low information content.

7.5
Creation of genotype networks and determining network metrics

We used in-house Python script and the Python package

igraph



¹⁹⁷


to generate directed genotype networks. These are graphs in which TFBS variants are nodes (vertices, genotypes), and variants that differ in a single nucleotide are connected by an edge. Each node of this network is associated with the corresponding DNA sequence and the associated regulation strength. Each edge is directed, i.e., it corresponds to a binding-score-increasing mutation, and points from a TFBS variant with lower regulation strength to a neighbor with higher regulation strength. We extracted the largest weakly connected subgraph (the“giant component”


⁶


) of the network, and used it for all further analyses. This giant component comprises the vast majority (99%) of sequenced genotypes for all of our TF landscapes. We used in-house Python and R scripts for all network analyses described below.

Epistasis:


Epistasis refers to non-additive interactions between two or more mutations. It can impose severe constraints on molecular evolution, because the mutations that are beneficial in one genetic background may be deleterious in another


²⁴


. Epistasis between two mutations can be classified as magnitude, simple sign, or reciprocal sign epistasis, depending on the sign (i.e., positive or negative) of the fitness effect of individual mutations and their combinations


²⁴


. In magnitude epistasis, the effect of a mutation on regulation strength varies depending on the genetic background but the sign of this effect (increasing or decreasing regulation strength) does not. Simple sign epistasis occurs if one single mutant has a lower regulation strength than both the wild type and the double mutant, while the other single mutant has a regulation strength that is intermediate to the wild type and double mutant. Reciprocal sign epistasis occurs when both mutations independently decrease regulation strength, but their combination increases regulation strength. The presence of reciprocal sign epistasis is a necessary condition for the existence of multiple peaks in an adaptive landscape


<sup>93

,

94</sup>


.

To determine the incidence of epistasis in our landscapes, we employed a method that involves identifying all “squares” in a genotype network with the

motifs

function from the

igraph

library in R. Each square consists of a “wild-type” sequence, a double-nucleotide mutant, and the corresponding two single mutants. We assessed epistasis for each square along a single axis by selecting the highest- regulation strength sequence as the double mutant. Our analysis placed each square into one of three categories: no sign epistasis, simple sign epistasis, and reciprocal sign epistasis. The no sign epistasis category included both magnitude epistasis and additivity (no epistasis) without differentiating between them, because neither affects the accessibility of landscape peaks through only binding-increasing mutations


<sup>11

,

108</sup>


. We determined the proportion of all squares that fell into each category (see

Supplementary Table S5

).

Peaks:


A peak is a genotype (TF binding site variant) whose neighbors all convey lower regulation strength than itself. Two peaks are connected if they are neighbors and convey the same regulation strength. We refer to the genotype with the highest regulation strength as the summit or global peak


<sup>6

,

198</sup>


.

Genotype connectivity:


To evaluate the sparsity of our networks and the likelihood of peak misclassification due to the incompleteness of our landscapes, we used a metric called “genotype connectivity.” This represents the number of immediate (1-mutant) neighbors of a given genotype for which our experiment produced repression strength data. A perfect experiment creating a complete landscape data set would provide repression data for each of our 4

⁸

genotype and all of its 24 immediate neighbors. In practice, however, data for some genotypes and some of their neighbors is unavailable. We quantified the “relative connectivity” of any one genotype as the ratio of the actual number of adjacent genotypes for which we have repression data to the theoretical maximum of 24 neighbors. This measure helps us to address questions about the sparsity of our landscape and its implications. Specifically, it allows us to determine if our sample is biased, with some regions or genotypes being more connected than others. This is particularly important for assessing whether peaks and high peaks are less connected than non-peaks, which could potentially lead to peak misassignments.

Accessible Paths:


In the context of our genotype networks, we call a mutational path accessible if the regulation strength increases with each mutational step


<sup>6

,

198</sup>


. We systematically enumerated all shortest accessible paths, distinguishing them by their path length, i.e., by the number of mutational steps in a path. Notably, there can be multiple alternative accessible paths with the same number of mutational steps from a single starting genotype to a peak.

Basins of attraction:


The basin of attraction of a peak comprises all TFBS variants from which accessible paths to the peak exist. We refer to the basin’s size as the number of variants in the basin. We determined basin sizes by exhaustive enumeration.

Overlap between basins.

The basins of attraction of different peaks may comprise overlapping sets of variants. To determine the overlap between two basins B

₁

and B

₂

, we used the Jaccard index

J



<sup>199

,

200</sup>


, which is equal to the size of the intersection between two sets of variants divided by the size of their union:

7.6
Generating randomly shuffled landscapes

To generate uncorrelated random landscapes for our three TF landscapes, we performed a random shuffling of regulation strength values from our genotypes, followed by landscape analysis to identify peaks. To this end, we employed an ad-hoc python script to randomly permute experimentally measured regulation strength values among all genotypes in each of our landscape. This procedure preserves the distribution of regulation strengths in each landscape, but also creates an uncorrelated random landscape


<sup>91

,

97</sup>


. Following random shuffling, we analyzed each shuffled landscape’s topography with the same methods we had applied to the original (unshuffled) landscape to identify its peaks. To ensure statistical robustness, we repeated the random shuffling and peak identification 10

³

times. We then compared the number of peaks between the 10

³

shuffled landscapes and the original (unshuffled) landscape.

7.7
Principal component analysis

In order to investigate which nucleotides in a TFBS are most important for changes in regulation strengths, we performed a principal component analysis for each landscape (

Supplementary Figures S19

-

S21

). To this end, we first one-hot-encoded our data. This encoding represents each categorical value (nucleotide at each position of a DNA string) as a binary vector of length 4. It thus converts an entire DNA string of length L into a 4xL binary matrix. We used this binary matrix to perform PCA with the R base function

prcomp

.

8.
Simulated adaptive walks

We simulated the adaptive evolution of a population on our landscapes by performing two different types of random walks, “greedy” adaptive random walks, and random walks based on Kimura’s model of fixation probabilities.

For all simulations, we assumed that only point mutations occur and that the time it takes for a point mutation to become fixed in a population is much shorter than the time it takes for a new mutation to appear that will eventually go to fixation. This scenario is also called the strong selection weak mutation (SSWM) scenario


<sup>16

,

99

–

101</sup>


. Under this scenario, evolving populations are monomorphic most of the time, i.e., all individuals have the same genotype. This scenario is realistic when the product of effective population size

N

and mutation rate

μ

is small (

Nμ

< 1), which is the case for

E. coli

(N=1.8

ξ10


⁸


, μ=2ξ

10

^-10

)


⁶⁷


. It allows us to model adaptive evolution as an adaptive random walk in our landscapes. Our simulations also account for mutation bias, which means that different types of mutation have a different probability of occurring in a population. We use mutation biases that were experimentally determined for

Escherichia coli



¹⁰⁴


.

In a greedy adaptive random walk, starting from any one genotype, only the mutational neighbor that conveys the largest fitness advantage is fixed in a population. We initiated one greedy random walk from each non-peak genotype and terminated the walk once a fitness peak was reached, i.e., once every neighbor of the current genotype had lower fitness than the genotype itself. Because every greedy random walk is deterministic in the absence of neutral mutations, it is sufficient to initiate one greedy random walk per starting genotype.

Another well-established model for random walks allows for genetic drift. It uses fixation probabilities computed by Kimura


<sup>98

,

201

,

202</sup>


i.e.,

f

_ij


= (1 –

e


<sup>-2

s</sup>


) / (1 –

e


<sup>-2

Ns</sup>


), where

f

_ij


is the probability of fixing mutation

j

in the background of genotype

i

,

N

is the effective population size, and

s

is the selection coefficient, i.e. the difference in fitness between genotypes

i

and

j



<sup>98

,

201</sup>


. For a given pair of genotypes, the only parameter of this model is the effective population size

N

, for which we explored values of

N

=10

⁸

,

N

=10

⁵

and

N

=10

²

. Generally, the smaller the population size is, the larger is the probability that neutral or deleterious mutations become fixed in a population.

For these “Kimura” adaptive walks, we chose 15,000 random starting genotypes for each of the three values of

N

, and simulated 1,000 random walks for each of them. At each step, we randomly picked a mutation

j

, generated a random number in the interval [0, 1], and considered the mutation to become fixed if the random number fell within the interval [0,

f

_ij


]. Computationally, we accelerated this process by precomputing all fixation probabilities for all genetic backgrounds, and then used the

python

function

numpy.choice

to generate a random sample from the multinomial distribution of fixation probabilities at each step


²⁰³


. Because of genetic drift, Kimura’s random walks do not necessarily terminate when they reach a fitness peak. For this reason, we simulated each such random walk for a maximum of 25 mutational steps.

9.
Randomized landscape null model for peak accessibility

To evaluate whether the accessibility of strong regulatory peaks observed in the empirical landscapes exceeds random expectations, we constructed randomized “null” landscapes and repeated evolutionary simulations on them under identical conditionsas for the empirical landscapes.

Specifically, for each transcription factor landscape, we generated 10

^$

independently shuffled landscapes landscapes see

Supplemtnary Methods 7.6

) by permuting repression-strength values across genotypes while preserving the network of genotypes and its mutational neighborhood structure. This procedure maintains the number of genotypes, their connectivity, and the distribution of repression-strength values, while removing correlations between genotype and phenotype. All other aspects of the analysis were kept identical to those used for the empirical landscapes.

For each randomized landscape, we simulated adaptive evolution using the same Kimura random-walk framework as for the empirical data. We assumed a population size of 10

⁸

individuals, initiated adaptive walks from the same number of starting genotypes, and used identical stopping criteria, fixation probabilities, and mutational bias parameters as described in

Supplementary Methods 8

(

“Simulated adaptive walks”)

.

For each randomized landscape, we computed the fraction of adaptive walks that reached a high regulatory peak. This yielded a null distribution of peak-accessibility values for each transcription factor. We tested the null hypothesis that the observed fraction from the corresponding empirical landscape is greater than the mean fraction derived from this null distribution using a one-sided Monte Carlo permutation test (𝑁 = 10

^$

randomized landscapes). For this test, we used Z-scores computed as the difference between the observed accessibility and the mean of the shuffled-landscape distribution, normalized by the standard deviation of that distribution.

10.
Incorporating experimental uncertainty into adaptive walks

Fluorescence-based sort-seq measurements are subject to experimental uncertainty arising from finite fluorescence bin resolution, variability in fluorescence measurements, and sequencing noise. To account for this uncertainty when defining fitness relationships between genotypes and identifying peaks in the landscape, we incorporated empirically estimated noise into pairwise repression strength comparisons.

For each genotype, we computed repression strength (

S

) as a continuous quantity derived from the distribution of sequencing reads across fluorescence bins (see Methods). We quantified experimental uncertainty in

S

by a genotype-specific noise parameter (

τ



_G


), calculated as the standard deviation of repression strength across three biological replicates. This parameter defines an uncertainty interval within which repression strength values cannot be reliably distinguished.

We compared the repression strengths of pairs of genotypes while explicitly accounting for their associated experimental uncertainty. Let

S



_A


and

S

_B


denote the repression strengths of genotypes

A

and

B

, with corresponding uncertainty estimates 𝜏


_A


and 𝜏


_B


. We considered genotype

A

to exhibit significantly stronger repression than genotype

B

if

In this case, we assigned a directed edge from

A

to

B

in the genotype network. Conversely, genotype

A

was considered to exhibit significantly weaker repression than genotype

B

if

in which case we assigned a directed edge from

B

to

A

. If neither condition was satisfied—i.e., if the uncertainty intervals of the two genotypes overlapped—we considered their repression strengths to be indistinguishable within experimental uncertainty. Such genotype pairs were treated as neutral and connected by two directed edges, one in each direction.

Applying these rules to all genotype pairs yielded the uncertainty-aware genotype network, denoted 𝒢


^S,τ


. For comparison, we also constructed a noise-free genotype network, denoted 𝒢


^S


, in which experimental uncertainty was ignored (

τ



_G


= 0) and pairwise comparisons were based solely on point estimates of repression strength

S

. In both networks, peaks were defined as genotypes without outgoing edges.

Supplementary tables

Supplementary figures

Data availability

Sequencing data has been deposited in the NCBI database under the BioProject accession code: PRJNA1162449. The data generated in this study have been deposited in the Zenodo public repository and are accessible via the following DOI: 10.5281/zenodo.13838265. All code used for processing data and plotting, as well as the final processed data, plasmid sequences, and primer sequences are available in our GitHub repository. The computer code generated in this study has been deposited in the Zenodo public repository and is accessible via the following DOI: 10.5281/zenodo.13838265.

Acknowledgements

We gratefully acknowledge financial support from the Swiss National Science Foundation grant 310030_208174. We also extend our thanks to the UZH University Priority Research Program in Evolutionary Biology, the UZH flow cytometry facility, and the Functional Genomics Center Zurich for their technical support. We are especially thankful to Andrei Papkou for his guidance with computational analysis and for engaging in theoretical discussions.

Additional information

Author contributions

C.A.W. and A.W conceived the study and designed experiments. C.A.W. carried out experiments. C.A.W and A.W. analyzed data. C.A.W wrote computer code to carry out bioinformatic work and analysis. C.A.W generated figures. L.G. wrote computer code to carry out simulations. C.A.W. and A.W. wrote the paper, which was edited by all authors.

Funding

Swiss National Science Foundation (310030_208174)

• 
  

  
  Cauã Antunes Antunes Westmann
  

  
  


• 
  

  
  Leander Goldbach
  

  
  

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