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Part Characterization

Bacteriophage lambda cI ([http://partsregistry.org/wiki/index.php?title=Part:BBa_K150004 BBa_K150004])

cI protein - an introduction

Bacteriophage λ cI exists as an inactive monomer at very low concentrations (<10^-9 M) but forms functional homodimers at physiological concentrations that remarkably lack a global symmetry [1]. It consists of 236 amino acids, although 237 amino acids are translated since the initiator methionine is removed by the host. Although λ cI is commonly called a repressor because of its negative regulatory functions at o_L and o_R, cI protein acts also as a positive regulator of gene transcription and can activate transcription of its own gene through the p_RM promoter in bacteriophage λ. In genetic and biochemical studies as well as through the crystal structure it has been shown, that the carboxy-terminal domain of the cI protein contains the major sites for dimerization and oligomerization [2, 3]. The cI protein has a two-domain structure with an N-terminal portion involved in DNA binding, and a C-terminal domain that mediates dimer formation, dimer-dimer interaction, and self-cleavage. The self-cleavage reaction is triggert when the lysogenic cell suffers DNA damage and depends upon an activated form of the bacterial RecA protein [2, 4-6]. The ‘hinge region’ connecting both domains contains a conserved site that can undergo this RecA-mediated autodigestion resulting in inactivation of the repressor by separating the N-terminal from the C-terminal domain of the repressor [4, 7]. The cI protein binds symmetrically to DNA, so that each amino-terminal domain contacts a similar set of bases. The N-terminal DNA-binding domain is made up of 5 α-helices of which helix 2 and helix 3 (the helix-turn-helix motif) are involved in nucleotide sequence specific DNA recognition and binding to the major groove of DNA [8-11]. [back]

Function of cI protein in bacteriophage λ

The cI repressor in bacterophage λ is the key component of a ‘genetic switch’ that enables the phage to transition from lysogenic growth to lytic development. The cI protein binds the o_L and o_R operator which overlap with the p_L and p_R promoter (the lytic promoters). This binding allows the maintenance of the lysogenic state to be governed by cI alone. As soon as cI is inactivated, e.g., by the SOS response after UV damage or other agents that cause DNA damage, the lytic development follows [12]. The two operators o_L and o_R both contain three binding sites for cI protein. In each case, site 1 (i.e. o_L1 and o_R1) has a ~10-fold greater affinity than the other sites for cI protein. The repressor, therefore, always binds to o_L1 and o_R1 first and than to the other sites in the operator in a cooperative manner [13, 14]. The carboxy-terminal domains of the repressor dimers mediate this cooperativity which improves the specificity and strength of the cI DNA binding, enabling strong repression of the lytic promoters [15]. Furthermore, another cooperative interaction between these two sets of tetramers bound to o_L and o_R, 2.4 kb apart, leads to the formation of a DNA loop held by a cI octamer, i.e. two interacting tetramers, that enhances repression of the early promoters [16-19]. In this DNA-multiprotein complex, the cI dimer bound at o_R2 represses p_R and at the same time also stimulates p_RM transcription, thus activating cI synthesis in a repressed prophage by a positive autoregulatory loop [20-22]. As the cI concentration increases because of p_RM activation, two additional cI dimers are recruited to bind o_L3 and o_R3 to further stabilize the o_L – o_R loop. In this context, cI overexpression is prevented by the binding of cI to o_R3 which represses p_RM [17, 19]. This positive and negative autoregulation at p_RM by cI ensures a narrow range of cI repressor level to be maintained, which is optimum for stable lysogeny but is at the same time adjusted low enough for efficient induction of the lysogen. By repressing transcription from the p_R promoter not only expression of genes in that operon is inhibited but also phage DNA replication by preventing transcriptional activation of λ ori, the site where phage DNA replication is initiated [23]. Even if O and P functions are present this inhibition occurs and appears to be critical for establishing a lysogen [24, 25]. The stable lysogen produces sufficient repressor not only to block prophage lytic development but also to block lytic development of any extraneous infection phage, thus imparting immunity to the lysogen against lytic superinfection [26]. [back]

Figure 1: λ cI repressor dimer bound to an operator fragment. A ribbon representation of the three-dimensional structure of the dimer complex crystallized by Stayrook et al. [1]. The N-terminal domain interacts with the operator DNA, the C-terminal domain self-associates and forms the dimer contacts. The two domains are connected by a linker that tethers the N-terminal domain to the C-terminal domain.

Figure 2: Structure of the λ cI C-terminal domain octamer. A ribbon representation of the three-dimensional structure of the octameric complex crystallized by Bell and Lewis [27]. Each of the four carboxy-terminal domain dimers that comprise the octamer are colored differently. The amino-terminal residues of each subunit are indicated. Because the amino-terminal residue of each subunit in the octamer points to the outside of the structure, the DNA-binding domains would fan around the structure. Since the structure closes up to itself, no higher-ordered oligomers can be formed.

Characterization of cI ([http://partsregistry.org/wiki/index.php?title=Part:BBa_K150004 BBa_K150004])

For the characterization of λ cI protein its ability to repress lytic development of bacteriophage λ was measured over a range of temperature from 28 °C to 42 °C in a phage burst experiment. If the used cI is functional no plaques should appear in a lawn of cells harboring this cI when infected with a phage that lacks its natural cI. In control cells in contrast every of this virulent particles should lead to a plaque.

Protocol

Preparation of plating bacteria

A overnight culture of the appropriate E. coli strain was grown in LB medium containing 10 mM MgSO₄ and 0.2 % maltose at 30 °C to reduce the amount of cell debris in the medium. For this characterization E. coli Top10 and E. coli Top10 with transformed cI were used. The added maltose leads to a substantial induction of the maltose operon including the lamb gene, which encodes the cell surface receptor to which bacteriophage λ binds. After harvesting the cells at 3000g for 10 minutes they were resuspended in 10 mM MgSO₄ and diluted to a final concentration of 2.0 OD₆₀₀. The suspension of plating bacteria was stored at 4 °C for up to 1 week.

Bacteriophage λ plaque assay

Tenfold serial dilutions of the bacteriophage λ stocks were prepared. In this case a bacteriophage mutant was used lacking a functional cI which would therefore always lead to plaques. From these dilutions 100 µl were mixed with the same amount of plating bacteria, incubated for 20 minutes at 37 °C to allow the bacteriophage particles to adsorb to the bacteria, this mixture than added to 3 ml molten top agar which was kept liquid at 48 °C and the entire contend poured onto a agar plate. After harden of the top agar the inverted plates were incubated at indicated temperatures over night. On the next day plaques could be counted.

Results

As it can be seen in figure 3 – 5 the cells harboring the part [http://partsregistry.org/wiki/index.php?title=Part:BBa_K150004 BBa_K150004] are not lysed by the used phage. This means that the constructed cI protein generator successfully expresses high levels of cI protein which is functional. Through the use of different temperatures it could be shown that the ability of the used cI protein to repress lytic development is not dependent of the temperature.

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Figure 3: phage burst experiment at 37 °C. Shown is the number of plaque forming units (pfu) over the dilution from our bacteriophage λ stock. E. coli Top10 that express cI protein (green curve) as well as untransformed E. coli (red curve) as control were infected with a bacteriophage λ lacking its own cI. After growth at 37 °C over night the amount of plaques was counted the next day. Note, that the used cI protein generator successfully blocks the phage burst.

Figure 4: phage burst experiment at 42 °C. Shown is the number of plaque forming units (pfu) over the dilution from our bacteriophage λ stock. E. coli Top10 that express cI protein (green curve) as well as untransformed E. coli (red curve) as control were infected with a bacteriophage λ lacking its own cI. After growth at 42 °C over night the amount of plaques was counted the next day. Note, that the used cI protein generator successfully blocks the phage burst.

Figure 5: phage burst experiment at 28 °C. Shown is the number of plaque forming units (pfu) over the dilution from our bacteriophage λ stock. E. coli Top10 that express cI protein (green curve) as well as untransformed E. coli (red curve) as control were infected with a bacteriophage λ lacking its own cI. After growth at 28 °C over night the amount of plaques was counted the next day. Note, that the used cI protein generator successfully blocks the phage burst.

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References

[1] S. Stayrook, P. Jaru-Ampornpan, J. Ni, A. Hochschild, M. Lewis, Crystal structure of the lambda repressor and a model for pairwise cooperative operator binding, Nature 452 (2008) 1022-1025.

[2] C.O. Pabo, R.T. Sauer, J.M. Sturtevant, M. Ptashne, The lambda repressor contains two domains, Proc Natl Acad Sci U S A 76 (1979) 1608-1612.

[3] C.E. Bell, P. Frescura, A. Hochschild, M. Lewis, Crystal structure of the lambda repressor C-terminal domain provides a model for cooperative operator binding, Cell 101 (2000) 801-811.

[4] J.W. Little, Autodigestion of lexA and phage lambda repressors, Proc Natl Acad Sci U S A 81 (1984) 1375-1379.

[5] R.T. Sauer, M.J. Ross, M. Ptashne, Cleavage of the lambda and P22 repressors by recA protein, J Biol Chem 257 (1982) 4458-4462.

[6] R.T. Sauer, C.O. Pabo, B.J. Meyer, M. Ptashne, K.C. Backman, Regulatory functions of the lambda repressor reside in the amino-terminal domain, Nature 279 (1979) 396-400.

[7] J.W. Little, LexA cleavage and other self-processing reactions, J Bacteriol 175 (1993) 4943-4950.

[8] C.O. Pabo, M. Lewis, The operator-binding domain of lambda repressor: structure and DNA recognition, Nature 298 (1982) 443-447.

[9] L.J. Beamer, C.O. Pabo, Refined 1.8 A crystal structure of the lambda repressor-operator complex, J Mol Biol 227 (1992) 177-196.

[10] A. Hochschild, Transcriptional activation. How lambda repressor talks to RNA polymerase, Curr Biol 4 (1994) 440-442.

[11] C.O. Pabo, R.T. Sauer, Transcription factors: structural families and principles of DNA recognition, Annu Rev Biochem 61 (1992) 1053-1095.

[12] A. Hochschild, The lambda switch: cI closes the gap in autoregulation, Curr Biol 12 (2002) R87-89.

[13] A.D. Johnson, B.J. Meyer, M. Ptashne, Interactions between DNA-bound repressors govern regulation by the lambda phage repressor, Proc Natl Acad Sci U S A 76 (1979) 5061-5065.

[14] M. Ptashne, Genetic switch: phage lambda revisited, 2nd ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, 2004.

[15] I.B. Dodd, K.E. Shearwin, J.B. Egan, Revisited gene regulation in bacteriophage lambda, Curr Opin Genet Dev 15 (2005) 145-152.

[16] I.B. Dodd, A.J. Perkins, D. Tsemitsidis, J.B. Egan, Octamerization of lambda CI repressor is needed for effective repression of P(RM) and efficient switching from lysogeny, Genes Dev 15 (2001) 3013-3022.

[17] I.B. Dodd, K.E. Shearwin, A.J. Perkins, T. Burr, A. Hochschild, J.B. Egan, Cooperativity in long-range gene regulation by the lambda CI repressor, Genes Dev 18 (2004) 344-354.

[18] B. Revet, B. von Wilcken-Bergmann, H. Bessert, A. Barker, B. Muller-Hill, Four dimers of lambda repressor bound to two suitably spaced pairs of lambda operators form octamers and DNA loops over large distances, Curr Biol 9 (1999) 151-154.

[19] S.L. Svenningsen, N. Costantino, D.L. Court, S. Adhya, On the role of Cro in lambda prophage induction, Proc Natl Acad Sci U S A 102 (2005) 4465-4469.

[20] B.J. Meyer, M. Ptashne, Gene regulation at the right operator (OR) of bacteriophage lambda. III. lambda repressor directly activates gene transcription, J Mol Biol 139 (1980) 195-205.

[21] D. Jain, B.E. Nickels, L. Sun, A. Hochschild, S.A. Darst, Structure of a ternary transcription activation complex, Mol Cell 13 (2004) 45-53.

[22] B.E. Nickels, S.L. Dove, K.S. Murakami, S.A. Darst, A. Hochschild, Protein-protein and protein-DNA interactions of sigma70 region 4 involved in transcription activation by lambdacI, J Mol Biol 324 (2002) 17-34.

[23] M.E. Furth, W.F. Dove, B.J. Meyer, Specificity determinants for bacteriophage lambda DNA replication. III. Activation of replication in lambda ric mutants by transcription outside of ori, J Mol Biol 154 (1982) 65-83.

[24] K. Mensa-Wilmot, K. Carroll, R. McMacken, Transcriptional activation of bacteriophage lambda DNA replication in vitro: regulatory role of histone-like protein HU of Escherichia coli, EMBO J 8 (1989) 2393-2402.

[25] M.S. Wold, J.B. Mallory, J.D. Roberts, J.H. LeBowitz, R. McMacken, Initiation of bacteriophage lambda DNA replication in vitro with purified lambda replication proteins, Proc Natl Acad Sci U S A 79 (1982) 6176-6180.

[26] D.L. Court, A.B. Oppenheim, S.L. Adhya, A new look at bacteriophage lambda genetic networks, J Bacteriol 189 (2007) 298-304.

[27] C.E. Bell, M. Lewis, Crystal structure of the lambda repressor C-terminal domain octamer, J Mol Biol 314 (2001) 1127-1136.

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oriT R ([http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003])

We have characterized the oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] in a conjugation system in which pUB307 is used as helper plasmid and E. coli as donor and recipient cells. We measured quantitatively the conjugative competence of oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] by conjugation between different strains, at different temperatures and with different donor/recipient ratios. We also compared the conjugation kinetics of [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] and pUB307 quantitatively.

The part oriT R [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] was cotransformed together with pUB307 (helper plasmid) into E. coli strain Top10. These cells therefore became conjugation donor and have a kanamycin (from pUB307) and ampicillin (from [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003]) resistance. The recipient cells have the plasmid pBAD33 with a chloramphenicol resistance. After mixing both cell types the conjugation starts and [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] as well as pUB307 can be transported. The helper plasmid pUB307 can still be transported since it still contains its oriT region. The transconjugants which have received [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] through the conjugation have ampicillin and chloramphenicol resistance and can therefore be easily selected on agar plates containing these two antibiotics. [back]

Test 1 Conjugation kinetics of oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] between E. coli Top10

On the first day we inoculated LB + antibiotics medium with a single colony or from a glycerol stock. For this test we used donor and the recipient cells that were both E. coli Top10. On the second day we washed the cells of the overnight culture twice with LB medium, transferred the donor cell suspension to the pellet of the recipient cells and resuspended the recipient cells in this medium. After another centrifugation step we resuspended the pellet which now is a mixture of donor and recipient cells in 100 µl LB medium without antibiotics. We than transferred the cell suspension on a membrane filter, which was placed on LB plates before and incubated these plates at 37 °C for 0, 6, 12, 18, 24, 30, 36, 42 minutes. Afterwards the membrane was put into 1 ml LB medium and the cells were resuspended by vortexing for 30 seconds. The cell suspension was diluted 10^-5 and plated on the LB / ampicillin + chloramphenicol (CA) plates. The plates were incubated at 37 °C overnight and on the third day the colonies could be counted.
We have done this test thrice independently and calculated the conjugation efficient which is defined as [transconjugants per ml] / [donor per ml] from this assay.
To count the donor and recipient cell density small aliquots of the used suspensions were diluted 10^-7 and plated on agar plates containing kanamycin + ampicillin for donor cells and chloramphenicol for the recipients. [back]

Table 1: Conjugation kinetics of oriT BBa_J01003 between E. coli Top10

Figure 1: conjugation kinetic of [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] between E. coli Top10 strains. Note that the conjugation efficiency increases with the time allowed for the conjugation to take place.

Figure 1 shows that the kinetic of the conjugative plasmid [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] has three phases. In the first (0 min to 24 min) and in the third phase (30 min to 42 min) the curve has a smaller slope, whereas the second phase (24 min to 30 min) it has a very big slope. An explanation for this result could be that the cell growth in the first 24 min can be neglected due to the average generation time of E. coli in the LB medium which lies between 20 and 30 minutes whereas in the second phase the growth of E. coli dominates the conjugation process and leads to this deviation from linearity. In this test we used very concentrated bacteria suspension in a small conjugation volume which leads to the observation of conjugation in an approximately linear process in the primary phase.
We have done a linear regression analysis for the first phase (0 min to 24 min) and we calculated the slope for this phase as 9.00 • 10^-3 [transconjugants • donor^-1 • minute ^-1]. We calculated the conjugation rate for [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] which is defined as transconjugants • donor^-1 • recipient^-1 • minute ^-1. The conjugation rate of [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] is 1.05 • 10^-12 [ml • min^-1 • cell^-1]. This rate was used by the modeling group as a parameter value.
Figure 1 also shows us that the transport of the plasmid pSB1A2 containing the part [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] by conjugation is a very fast process since we got transconjugants in the first minutes of the experiment. Through this experiment we can conclude that the transport of the plasmid pSB1A2 containing [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] by conjugation needs only about 2 minutes. Since we have two conjugative plasmids in our donor cells, we wanted to see which plasmid is preferred to be transported by conjugation. We therefore plated the cells in the test C (see table 2) also on agar plates containing chloramphenicol + kanamycin (CK) and chloramphenicol + kanamycin + ampicillin (CKA) to select for transconjugants getting pUB307 and both plasmids. [back]

Table 2: Test C conjugation kinetics of oriT BBa_J01003 and pUB307 between E. coli Top10

Figure 2: Conjugation efficiency of different plasmids. Transconjugants with [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] were selected by plating on LB agar containing chloramphenicol and ampicillin, transconjugants with pUB307 by plates containing chloramphenicol and kanamycin, and transconjugants with both plasmids by plates containing chloramphenicol and kannamycin and ampicillin.

Figure 2 clearly shows that the plasmid pSB1A2 containing only [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] is preferably transported during the first 20 minutes in comparison to pUB307. Since pUB307 has the same oriT as [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] is, we assume the size of the plasmid to be the reason for this since [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] in pSB1A2 is approx. 2.5 kb in size whereas pUB307 has 60 kb. A comparison of the red and the green curve shows us that the difference in the duration of the transport of plasmids with different size is not very big. Looking at the blue curve you can see that the first transconjugants with both plasmids can be detected after approx. 18 minutes which is longer than the sum of both transport times. [back]

Test 2 Conjugation kinetics of oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] between different E.coli strains

For this experiment we used the same technique as described in test 1. In this case we used different E. coli strains as recipients, namely E. coli Top10, MG1655 and DH5α. In this experiment the cells on the membranes were incubated for 0, 6, 12, 18, 24, 30, 36 and 42 minutes at 37 °C. This test was conducted twice independently and the conjugation efficiency calculated as described in test 1. [back]

Table 3: Conjugation kinetics of oriT BBa_J01004 between different E. coli strains

Figure 3: Conjugation of [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] in pSB1A2 between different strains of E. coli. The transconjugants were selected through growing on plates containing chloramphenicol and ampicillin. Note that conjugation from Top10 to Top10 and DH5α is comparable whereas conjugation in MG1655 takes place a lot slower.

As you can see in figure 3 there is a difference in the conjugation kinetic between Top10 as donor and different strains as recipient. With E. coli Top10 as donor E. coli DH5α is a comparably good recipient as Top10 whereas MG1655 showed up as a bad recipient under these conditions. Comparing the red with the blue curve one can see that DH5α transconjugants appear later (6 minutes) that Top10, but the conjugation efficiency and also the conjugation rate (slope between 6 and 24 minutes) are in the same range with Top10 and DH5α. We assume that more time is needed for stabilizing the pillus contact between Top10 and DH5α than between two Top10 cells, but the conjugation between Top10 and DH5α can be done as good as between two Top10 cells. Comparing the red and the green curve one can see that the first MG1655 transconjugants appear much later (20 minutes) and the conjugation efficiency and rate (slope between 20 and 32 minutes) are much smaller. Therefore we conclude that conjugation between different strains is possible using this oriT but the efficiency depends on the used strains and their compatibility. [back]

Test 3 Conjugative competence of oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] at different donor/recipient ratios

In this experiment we again used the same protocol as described above in test 1 but used different donor / recipient rations, namely 50:1, 25:1, 10:1, 8:1, 5:1, 3:1, 2:1, 1:1, 1:2, 1:3, 1:5, 1:10, 1:25, and 1:50. This test was conducted twice independently and the conjugation efficiency calculated as described above. The donor and recipient cell density was measured by plating 10^-7 dilutions on agar plates containing kanamycin + ampicillin for the donor and chloramphenicol for recipient cells. [back]

Table 4: Conjugative competence of otiT BBa_J01003 at different donor / recipient ratios

Figure 4: Conjugative competence of oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] at different donor / recipient ratios. For all of the samples a similar cell density was used. For the conjugation a time of 20 minutes was set. The transconjugants were selected by plating on plates containing chloramphenicol and ampicillin.

Figure 4 shows very clearly that at same cell density, if donor : recipient ratio is 1 : 2 the highest amount of transconjugants can be achieved. A big difference in the amount of donor : recipient leads to a low number of transconjugants. At low donor : recipient ratios the conjugation efficiency was quiet high but lead to a low number of transconjugants since the density of donor cells was too low. Only at donor : recipient rations between 1 : 1 and 1 : 3, a high conjugation efficiency and a high donor density was achieved leading to a high number of transconjugants. A maximum in the number of transconjugants could be achieved at a ratio of 1 : 2. [back]

Test 4 Conjugative competence of oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] at different temperatures

In this experiment the same method is used as described above in test 1. Here, donor and recipient both were E. coli Top10. The incubation during conjugation took place at different temperatures, namely 4 °C, 22 °C, 30 °C, 37 °C and 42 °C for 20, 40 and 60 minutes. For counting of transconjugants the cell suspension was diluted 10^-6 and plated on agar containing ampicillin and chloramphenicol (CA). This experiment was conducted twice independently. The donor and recipient cell density was again calculated by plating 10^-7 dilutions on plates containing kanamycin and ampicillin or chloramphenicol, respectively. [back]

Table 5: Conjugative competence of otiT BBa_J01003 at different temperatures

Figure 5: Conjugative competence of oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] at different temperatures. All of the samples have the similar cell density and similar donor : recipient ratio. The conjugation time was set to 20, 40 and 60 minutes. Transconjugants are selected by plating on the agar containing chloramphenicol and ampicillin. Note, that the conjugation efficiency after 20 and 40 minutes is the same at 37 °C and 42 °C but differs after 60 minutes.

As you can see in figure 5, conjugation is most efficient at 37 °C. From 0 to 20 minutes during the primary conjugation phase the cell growth can be neglected. From 20 to 40 minutes, the cell growth is also presented in the conjugation curves. At 4 °C, the conjugation curve has a very small slope (near zero) and no significant transconjugants production could be observed. At 22, 30, 37 and 42 °C, a significant number of transconjugants could be achieved, but the conjugation efficiency and the conjugation rate (slope between 0 and 20 minutes) differ strongly. From 22 °C to 37°C, the conjugation efficiency and conjugation rate increases with rising temperature. Comparing these three curves between 20 and 40 minutes one can see that the cell growth gets more important with rising temperature. Dynamically, the cells have more mobility at higher temperature, and have a bigger possibility to touch another cell, which is necessary for conjugation. When we compared the conjugation curve at 42 °C with the one at 37 °C, one can see, that the conjugation efficient and the conjugation rate at 42 °C is as good as at 37 °C. After the primary conjugation phase, the production of transconjugants at 42 °C is clearly worse than the production at 37 °C what can be seen at comparing the curves between 40 and 60 minutes. A possible reason for this could be a lower growth and higher death rate at 42 °C compared to 37 °C. So, one can conclude, that for best conjugation efficiency one needs a warm environment for the conjugation to take place but also temperatures which favor cell growth. [back]

Summary

The characterization of [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] in pSB1A2 has given the following results:

• The conjugation has an average conjugation rate of 1.05 • 10^-12 [ml • min^-1 • cell^-1]
• The conjugation has an approximate transport time of 2 minutes
• The oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] will be transported first in attendance of pUB307 in the donor cells
• Smaller plasmids favor the conjugation and are conjugated first
• The oriT [http://partsregistry.org/wiki/index.php?title=Part:BBa_J01003 BBa_J01003] can be conjugated between Top10 cells and from Top10 to DH5α cells with similar efficiency but from Top10 to MG1655 with a strongly lower efficiency
• The conjugation favors a donor : recipient ratio of 1 : 2
• The conjugation has the best production of transconjugants, the best efficiency and the best conjugation rate at 37 °C

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Chloramphenicol resistance cassette (BBa_K150003)

Figure 2.1

Figure 2.1

Figure 2.1

LuxR-Colicin E1-Receiver ([http://partsregistry.org/wiki/index.php?title=Part:BBa_K150009 BBa_K150009])

ColicinE1 Producer Controlled by 3OC6HSL Receiver Device - Introduction

The constitutive expressed LuxR protein is able to build a complex with an AHL molecule. [7] This complex functions as transcription factor for the lux pR promoter and activates therefore the colicinE1 production. The activation of the lux pR promoter leads additionally to expression of the kil protein, also called lysis protein by read through of a terminator located in the colicinE1 operon. Production of the lyses protein leads to lysis of the host cell and colicin release. The colicinE1 immunity protein (imm) located on the opposite strand is expressed under a constitutive promoter and protects the host cell from lysis up to a certain threshold of the lysis protein. [back]

Colicin E1 - Mode of Action

Colicin E1 is a 522 residue containing protein and belongs to the group A colicins. It kills sensitive cells, which harbor no ColE1 plasmid but contain the appropriated receptor BtuB, by forming pores, or more precisely ion channels into the inner membrane. [1] Affected cells can be E. colis or related strains but also some eukaryotic cells have been affected by the toxic activity of colicin. [2] The first step of the lethal action of colicin E1 is binding to the vitamin B12 receptor (BtuB). Colicins are able to kill bacteria by a “one hit” mechanism. This works due to destruction of the target cells cellular energy. The voltage dependant channel in the cytoplasmatic membrane leads to a depolarization of the target cell. The conductance of the channel has been shown to be ~10⁷ ions/channel-sec in 1 M NaCl. [3] Its ability to form pores is given by the C-terminal domain. This domain consists of 10 tightly packed α–helices (see Figure 1). [4], [6] The colicin protein is a water soluble molecule containing mostly amphipathic helices except from two hydrophobic ones. Due to these two hydrophobic helices the protein is able to target the cell membrane and act as a membrane protein. [back]

Figure 1 (a) Ribbon diagram of the Colicin E1 pore-forming domain by X ray crystal structure. The channel forming domain contains 10 helices (I-X). The two hydrophobic helices (VIII and IX) are labeled yellow. (b) Colicin E1 in a membrane bound state in two perspectives. Left panel shows the surface bound helices with an area of ~4200 Å2. Right panel shows colicin E1 bound to a lipid bilayer. The intra-membrane sections of the two helices VIII and IX are shown in yellow.

Killing efficiency on bacteria

To measure the killing efficiency and which amount of cells or colicins are needed to reach any killing activity a colicin activity test was carried out. Therefore bacteria containing the colicin part ([http://partsregistry.org/wiki/index.php?title=Part:BBa_K150009 BBa_K150009] in TOP10 or MG1655) and GFP producing cells (reference promoter, TOP10) were inoculated in TB-media with appropriated antibiotics at 37 °C for 4 to 6 hours and the optical density of the two strains was adapted. The colicin cells were added in different ratios to a constant amount of GFP producing cells. The total volume was kept constant and the missing amount added with TB-media (no antibiotics). The colicin production was induced by several concentrations (0 M-100 nM) of N-Acyl-Homoserin-Lactone (AHL). The OD and GFP intensities were measured at 37 °C in the Tecan Microplate Reader every half an hour for about 12 hours. As a negative control the similar test was carried out with cells, containing the same plasmid without the colicin gene on it. Figure XX2 shows results of tests with a prey-killer ratio of 100:1. Due to the correlation of the GFP intensity and the optical density, the GFP intensity was used as marker for the living prey cells. All experiments using killer cells and high AHL concentrations in the medium all prey cells were killed. In reference experiments (see Figure 2, bottom) using E. coli TOP 10 cells harboring a LuxR-receiver without colicin operon (comparable to part [http://partsregistry.org/wiki/index.php?title=Part:BBa_T9002 BBa_T9002] without GFP), the prey cells were able to grow for each AHL concentration. Consequently the lethal action of the killing part could be shown. [back]

Figure 2: Results of colicin E1 toxicity tests for different AHL concentrations with a prey-killer ratio of 100:1. Top left panel: GFP intensities of prey cells treated with killer cells over the time. Top right panel: Optical densities of prey cells treated with killer cells plotted against the time. Bottom left panel: GFP intensities of prey cells treated with reference cells versus time. Top right panel: Optical densities of prey cells treated with reference cells over the time. The graphs proof the toxicity of colicin E1 as well as the functionality of the part [http://partsregistry.org/wiki/index.php?title=Part:BBa_K150009 BBa_K150009]. Each killer cell is able to kill up to 100 prey cell if there is a sufficient amount of AHL present.

Additionally it can be seen that the killing efficiency is related to the AHL concentration in the medium. Figure 1Figure 3 shows a dose-response curve of GFP intensity after 12 hours dependent on the AHL concentration. The P_LuxR promoter has to be activated with an AHL concentration higher than 500 pM. Thus enough colicin E1 is produced and released to kill the prey cells (see Figure 3). [back]

Figure 3: Dose-response curve of AHL concentration and killing efficiency of the killer strain for a prey-killer ratio of 100:1. The GFP intensities at t = 12 h are plotted against AHL concentrations (0 M – 1 nM). For AHL concentrations higher than 600 nM all preys were killed.

Regarding the prey survival, dependent on the prey-killer ratio, several effects can be observed (see Figure 4). Having a prey-killer ratio of 1:1, or even a higher killer fraction, all prey cells were already killed when no AHL was present. This effect is caused by the leakiness of the P_LuxR promoter. For prey-killer ratios between 5:1 and 100:1 the killing efficiency can be regulated by the AHL concentration. In experiments having prey-killer ratios higher than 100:1 the growth of the prey strain was not influenced. [back]

Figure 4: Toxicity of LuxR-colicin E1-receiver part for different prey-killer ratios. The graphs show GFP intensities of prey cells at t = 0 h and t = 12 h. For prey-killer ratios of 1:1 all prey cellss were killed after 12 hours even if there was not AHL present. For prey-killer ratios between 5:1 (not shown) and 100:1 the killing efficiency depends on the AHL concentration in the media. If there is a greater predominace of prey cells (500:1) the growth of the prey population is not influenced.

Prey killer system

To measure the functionality of the killer-prey system were tested. Therefore bacteria containing the colicin E1 part ([http://partsregistry.org/wiki/index.php?title=Part:BBa_K150009 BBa_K150009] in TOP10 or MG1655) and prey cells containing an AHL-producing part ([http://partsregistry.org/wiki/index.php?title=Part:BBa_K150000 BBa_K150000]) were inoculated in TB-media with appropriated antibiotics at 37 °C for 4 to 6 hours. The optical density of the two strains was adapted. The colicin E1 producing cells were added in different ratios to a constant amount of prey cells. The total volume was kept constant and the missing amount added with TB-media (no antibiotics). The colicin production was induced by several concentrations (0 M-100 nM) of N-Acyl-Homoserin-Lactone (AHL). The OD and GFP intensities were measured at 37 °C in the Tecan Microplate Reader every half an hour for about 12 hours. As a negative control the similar test was carried out with cells, containing the same plasmid without the colicin E1 gene on it. Figure 5 shows that the prey cells were killed for prey-killer ratios from 1:1 up to 25:1. Therefore it is proven that our system works as expected. [back]

Figure 5: Prey-Killer system test. GFP intensity of prey strains was measured over 12 h at 37 °C. Prey cells were killed for prey-killer ratios from 1:1 up to 25:1 (blue, green, red, light blue) due to the colicin E1 production of the killer strain. This production was activated by the AHL secretion by the prey strain. In a reference experiment prey cells were able to grow (violet line).

Lysis of killing strain

Besides the killing efficiency the time frame of the lysis of the killer strain was analyzed. Therefore growth curves of killer cells ([http://partsregistry.org/wiki/index.php?title=Part:BBa_K150009 BBa_K150009] in E. coli TOP10 cells) induced with different AHL concentrations were measured (see Figure 6).

Figure 6: Lysis test with LuxR-ColicinE1-Receiver dependent on AHL concentrations. On the left panel the growth of a killer population induced with different concentrations of AHL is shown. At the right panel the data of referent experiments are plotted. For AHL concentrations between 0 M and 1 nM no influence on the growth of the killer population is observed. For concentrations between 5 nM and 25 nM the population grew in the first hour, afterwards a lysis effect can be observed

During the first hour of the measurement no effect could be observed. Afterwards cell lysis dependent on AHL concentrations was observed. For AHL concentrations between 0 M and 1 nM only few cells lysed. Thus growth of the population is hardly influenced. Concentrations from 5 nM up to 25 nM show a strong effect on the amount of lysed killer cells. For 5 nM and 10 nM the growth curves flatten out. After reaching a maximum during the third and fourth hour the population size decreases and converges to a constant value. For higher AHL concentrations, e.g. 25 nM, a similar effect can be observed, but much stronger. The population only shows a weak growing tendency but then converges directly to a constant size. In addition to these growth measurements lysis of the killer cells was observed under the microscope. Figure 7 shows killer cell which were induced by AHL (left panel, t = 0 min). After 30 minutes one cell is lysed (right panel). [back]

Figure 7: Visualization of the lysis effect of the killer cells. The prey (GFP) and the killer (mCherry) strain are mixed on a 0.5 % agarosepad. Killer cells were induced with AHL before adding to the medium. The killer cell on the left picture (white arrow) is lysed after 30 minutes. This effect is caused by the AHL induction. Colicin production and at same time in lower amounts the lyses protein production are induced leading to a lysis of the host cell and colicin release.

References

[1] Cramer W. et al. (1990) Structure and dynamics of the colicin E1 channel. Molecular Microbiology 4: 519-526.
[2] Smarda J. et al. (2001) Cytotoxic effects of colicins E1 and E3 on v-myb-transformed chicken monoblasts. Folia Microbiol. (Praha) 47: 11–13.
[3] Bullock J. et al. (1983) Comparison of the macroscopic and single channel conductance properties of colicin E1 and its COOH-terminal tryptic peptide. J. Biol. Chem. 258: 9908–9912.
[4] Elkins P. et al. (1997) A mechanism for toxin insertion into membranes is suggested by the crystal structure of the channel-forming domains of Colicin E1. Structure. 5: 443-458.
[5] Lindeberg M. et al. (2000) Unfolding Pathway of the Colicin E1 Channel Protein on a Membrane Surface. J. Mol. Biol. 295: 679-692
[6] Davies, J. et al. (1975) Genetics of resistance to colicins in Escherichia coli K12: cross-resistance among resistance of group A. J. Bacteriol. 123:102–117 [7] Engebrecht J. et al. (1984) Identification of genes and gene products necessary for bacterial bioluminescence. Proc. Natl. Acad. Sci. USA 81: 4154–58

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LuxI sender device ([http://partsregistry.org/wiki/index.php?title=Part:BBa_K150000 BBa_K150000])

General Description

AHL, a cell-cell signaling molecule is produced by the LuxI protein. The LuxI protein is expressed due to the luxI gene which is under the control of a constitutive promoter ([http://partsregistry.org/wiki/index.php?title=Part:BBa_J23107 BBa_J23107]). For this reason no input is needed for induction of gene expression. As output AHL can be measured. [back]

Quantification of AHL production

For AHL quantification the amount of AHL in supernatant of producing cells (E. coli TOP 10 containing [http://partsregistry.org/wiki/index.php?title=Part:BBa_K150000 BBa_K150000]) was determined. Therefore eight probes with TB-media were inoculated with 2 µl/ml of an AHL producing cell overnight culture at 37 °C. Sterile filtered supernatant of these probes was produced hourly for 7 hours and stored at 4 °C. To estimate the amount of AHL producing cells the optical density was measured before. (Figure 1, top left). [back]

Figure 1: Results of sender activity test. Top left: Growth curve of sender culture. Top right: GFP/OD values of sender supernatants at t = 10 h. Bottom left: GFP/OD – AHL calibration curve. Dotted line visualized regression of an exponential growth curve. Bottom right: Produced AHL concentrations of the sender cells.

For the AHL determination 1 ml supernatant was diluted with 1 ml TB-media and inoculated with a constant amount of AHL inducible cells which produce GFP after induction ([http://partsregistry.org/wiki/index.php?title=Part:BBa_T9002 BBa_T9002] in E. coli TOP 10). The OD and GFP intensities were measured at 37 °C half-hourly for about 28 hours. For calibration the same test were performed with distinct AHL concentrations from 0M to 50 nM (SIGMA). Figure 2 shows the GFP/OD development over the time depending on the sender supernatants (left panel) or the preassigned AHL concentrations (right panel). The values increase and reach a maximum after 6 hours. Then they decrease slightly. [back]

Figure 2: GFP/OD results of sender activity measurements. On The left panel the GFP/OD values for induction with sender supernatants are shown and on the right panel the GFP/OD values for predefined AHL inductions can be observed. During the first sic hours the values increase and reach a maximum until the decrease slightly. The values at t = 10 h were used for calculations of AHL concentrations in supernatants.

A GFP/OD – AHL calibration curve (Figure 1 bottom left) was calculated using the GFP/OD values at t = 10 h. An exponential growth curve was fitted to these data (blue dotted line; y = 4325 * e^0.00346*x). The AHL concentrations of the sender supernatants were determined using the GFP/OD values at t = 10 h and the exponential regression. In the course of the time the AHL concentration increase until it reaches a certain maximum before decrease slightly (see Figure 1, bottom right, Table 1). The maximum is reached at the end of the logarithmic phase. Several tests have shown that the maximal concentration that can be produced by part [http://partsregistry.org/wiki/index.php?title=Part:BBa_K150000 BBa_K150000], is approximately 1100 and 1300 pM.

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LuxS from Vibrio harveyi (partly characterized)

LuxS encodes for the protein (LuxS autoinducer synthase) that is responsible for the synthesis of autoinducer 2 (AI-2) in many species of Gram-negative and Gram-positive bacteria. This LuxS construct is cloned from Vibrio harveyi, a Gram-negative bioluminescent marine bacterium, which regulates light production in response to two distinct autoinducers (AI-1 and AI-2). [1] [back]

Functional check of LuxS/AI-2 production

The functionality of the new LuxS part that was cloned by PCR from V. harveyi (BB120) in a ptrc99α plasmid was proven by measurement of luciferase activity in Vibrio as a reporter strain. DH5α cells with the new construct pTrc99α_LuxS were grown up and the supernatant was harvested by using centrifugation and sterile filtration. The gained supernatant was added to the reporter cells in different dilutions and incubated for three hours. Luminescence and OD were measured after 0.5h and 3.5h. [back]

Results

Cell growth rate of all samples LuxS and Blank was shown to be similar in OD measurements (data not shown). Luminescence of Vibrio reporter strain (data shown in figure 1) seems to increase with the concentration of supernatant in the cell delusion. Blanks with supernatant of DH5α cells transformed with pTrc99α plasmid show a significantly smaller Luminescence and there is no increase of Luminescence for increasing supernatant concentration in the cell delusion. [back]

Figure 4: Vibrio cell delusions with different AI-2 supernatants 1:3, 1:5 and 1:10 were measured for luciferase activity after 30min and 3.5h incubation at 30°C using the Infinite® 200 microplate reader by Tecan, Switzerland; Supernatant sterile filtrated, no NaCl added (Protocol)

Reference

[1] The LuxS family of bacterial autoinducers: biosynthesis of a novel quorum-sensing signal molecule. Mol Microbiol. 2001 Jul; 41(2):463-76.

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