Advertisement

International Microbiology

, Volume 21, Issue 1–2, pp 15–22 | Cite as

The sequences of MinE responsible for its subcellular localization analyzed by competitive binding method in Escherichia coli

  • Miguel Á. Pérez-Rodríguez
  • Isabel Cristina Rodríguez-Luna
  • Ricardo Carreño-López
  • Edgar E. Lara-Ramírez
  • Mario A. Rodríguez-Pérez
  • Xianwu Guo
Original Article
  • 18 Downloads

Abstract

The subcellular localization of a protein is important for its proper function. Escherichia coli MinE is a small protein with clear subcellular localization, which provides a good model to study protein localization mechanism. In the present study, a series of recombinant minEs truncated in one end or in the middle regions, fused with egfp, was constructed, and these recombinant proteins could compete to function with the chromosomal MinE. Our results showed that the sequences related to the subcellular localization of MinE span several functional domains, demonstrating that MinE positioning in cells depends on multiple factors. The eGFP fusions with some truncated MinE from N-terminal resulted in different cell phenotypes and localization features, implying that these fusions can interfere chromosomal MinE’s function, similar to MinE36–88 phenotype in the previous report. The amino acid in the region (32–48) is sensitive to change MinE conformation and influence its dimerization. Some truncated protein structure could be unstable. Thus, the MinE localization is prerequisite for its proper anti-MinCD function and some new features of MinE were demonstrated. This approach can be extended for subcellular localization research for other essential proteins.

Keywords

MinE Subcellular localization Cell division Sequence 

Introduction

The subcellular localization of proteins is a basic issue in cell biology. Bacterium is a highly organized unicellular microorganism, wherein each protein takes up its specific place and functions at a given time so that many proteins in bacteria display conspicuous characteristics of localization. In essence, a protein in place means that the protein is anchored to other structural molecules of cells via interaction, called anchor proteins (Rudner and Losick 2010; Shapiro et al. 2009). Further, if some kinds of proteins show the same mechanism for interaction, they could share similar amino acid sequence, as the signal peptides in sorting eukaryotic proteins, to form a similar tridimensional structure (Emanuelsson et al. 2000). Hitherto, this issue was paid little attention.

In Escherichia coli, the spatial distribution of FtsZ ring (Z-ring) in cell is regulated by the Min system (MinCDE). MinD attaches to membrane only in ATP-bound form, locating at the cell pole. MinC, when captured and activated by MinD, has the inhibitor activity that prevents the Z-ring formation, resulting in MinCD complex attaching in the cell membrane. But MinE prevents the MinCD formation in the cell and forms an E-ring near the cell pole that promotes the release of MinD from the membrane. The two opposite processes result in pole to pole MinCD complex oscillations, consequently the highest MinCD concentration in the poles and the lowest concentration in the middle of cell, allowing the Z-ring formation (Hu and Lutkenhaus 2000; Zhou et al. 2005).

Thus, MinE is required for the right placement of Z-ring during the cell division in E. coli (de Boer et al. 1989; Pazos et al. 2014; Bramkamp and van Baarle 2009). MinE has three functional regions: the membrane-binding domain (residues 2–12), the anti-MinCD domain (residues 13 to 31), and the topological specific domain (TSD), also called dimerization domain (residues 32–88) (Shih and Zheng 2013). It forms homodimers that showed different function from monomer (King et al. 1999, 2000). MinE showed clear dynamic localization, oscillating from end to end of the cell in a MinD-dependent way (Hale et al. 2001; Hsieh et al. 2010).

To analyze the factors that determine the protein localization, an entry could be to study the protein sequence related to its localization information because the positioning of a protein in cells should be relevant to its amino acid sequence. Therefore, it is necessary to know which parts of sequences in a protein would affect its subcellular localization. The protein MinE was chosen as a model protein for this study. It is a small protein with only 88 amino acids (Zhao et al. 1995; Pichoff et al. 1995). It has been known some functions and interactions, which could help us to analyze the experimental results on localization. The rationale for the experimental design for a localized protein is based on the following hypotheses. (1) The expression of extrachromosomal full or truncated MinE fused to the eGFP can compete to bind other proteins or substrates with the chromosomal MinE. (2) If the sequence related to MinE positioning is present, the MinE should keep its subcellular localization in cells. (3) If the truncated MinE fusion on the plasmid shows its normal function as the chromosomal MinE, the fusions are able to compete with the chromosomal MinE and the cells show the phenotype similar to that of gene overexpression, whereas if fusions express with incomplete function, the fusions are able to interfere with the full function of chromosomal MinE protein and the cells will show similar phenotype as absence, less expression, or partial function of this gene. (4) If the recombinant MinE lost the ability to affect the chromosomal one, the recombinant also loses the localization feature. In the present study, we constructed a series of consecutively truncated minE fused to egfp (Cormack et al. 1996) under the constitutive promoter Pfla for expression, with a protocol based the In-Fusion® Dry-Down PCR Cloning (clonthech) without introduction of any restriction site. Expression of MinE fusion proteins caused the subcellular localization changes and cell form modification in E. coli under the presence of chromosomal MinE. The sequences that are involved in MinE subcellular localization and the cell shape preservation were then determined. Meanwhile, this simple method can generally be used for the protein localization sequence analysis.

Materials and methods

Strains, plasmids, and growth conditions for bacterial culture

E. coli DH5 alpha strain was grown in LB medium or LB medium supplemented with ampicillin (100 μg/ml) if used for transformation with plasmids. The Pfla-egfp segment was obtained from pDH80 vector (Cormack et al. 1996) and pBluescript SK-(Stratagene) vector as a carrier. The purification of plasmids was performed using the Wizard™ kit plus SV minipreps DNA purification (Promega). The genomic DNA was purified by Wizard™ Genomic DNA purification kit following manufacturer instructions.

DNA manipulation and plasmid construction

All the primers used in this study are listed in Table 1. Primers 1 and 2 were used to amplify the Pfla/egfp segment from pDH80 vector. The Pfla/egfp amplicon was cloned in pBluescript using XhoI-KpnI digestion to generate the pBS80 (Fig. S1A). pBS80 was then amplified by inverted PCR (rPCR) using the primer pairs 3–4 and 5–6 to generate two different linearized pBS80 (Fig. S1B). The pairs of primers 7–8 and 9–10 were added 15 nucleotides in the 5′ end, identical to the ends of linearized pBS80s, and used to amplify the MinE gene (Fig. S1C). Those linearized pBS80 and amplified minE were ligated by homologous recombination using the In-Fusion® Dry-Down PCR Cloning Kit, resulting in the plasmid pGM or pMG, which depends on the ligation as egfp-minE or minE-egfp, respectively (Fig. S1D). The pFGM came from the pGM lacking the lac promotor sequence (Plac). The pGM was amplified by rPCR using the primers DpL1 y DpL2, which shared 15 homolog nucleotides at the 5′ end. The pFGM was obtained by self-ligation of pGM-rPCR product using the In-Fusion® Dry-Down PCR Cloning Kit.
Table 1

Oligonucleotide sequences used in this study

Oligos

Sequences (5′-3′)

1

CGGCTCGAGATAAAGCCCTTTAAAATTTC

2

CGATGGTACCTTATTTGTATAGTTCATCCATGCC

3

ATGAGTAAAGGAGAAGAACTTTT

4

ATGTATATCTCCTTCTTAAATCTAGATT

5

GGTACCATCGAATCACTAGTGCGG

6

TTTGTATAGTTCATCCATGCCATGTGTAATCC

7

GAAGGAGATATACATATGGCATTACTCGATTTCT

8

TTCTCCTTTACTCATTTTCAGCTCTTCTGCTTC

9

GATGAACTATACAAAATGGCATTACTCGATTTCTTTCTCTCG

10

TGATTCGATGGTACCTTATTTCAGCTCTTCTGCTTCCGGT

DPL1

CTGGGGTGCCTAATGAGTGAGC

DPL2

CATTAGGCACCCCAGGAACAAAGCTGGGAGCTCCACC

F1

GGTACCATCGAATCACTAGTGCGG

1–1

TGATTCGATGGTACCTTAGTTCAGCTCAAGAATAGAAATATCGCCATCT

1–2

TGATTCGATGGTACCTTACTCAGGATCAATTTGTACGTATTTACAAATGACC

1–3

TGATTCGATGGTACCTTAATCTTTACGCAACTGCGGCAGATA

1–4

TGATTCGATGGTACCTTATGCATCGCTGCGACGGCGTTCAGCA

1–5

TGATTCGATGGTACCTTAAATAATCTGCAGCCGTTCTTTTGCAAT

1–6

TGATTCGATGGTACCTTAGTTGGCTGTGTTTTTCTTCCGCGA

F2

TGCCATTTTGTATAGTTCATCCATGCCA

2–1

CTATACAAAATGGCAAAGAAAAACACAGCCAACATTGCAAAAGAA

2–2

CTATACAAAATGGCAATTGCAAAAGAACGGCTGCAGATTAT

2–3

CTATACAAAATGGCACTGCAGATTATTGTTGCTGAACGCC

2–4

CTATACAAAATGGCAGAGGTCATTTGTAAATACGTACAAATTGATCCT

2–5

CTATACAAAATGGCACAAATTGATCCTGAGATGGTAACCGTACAG

2–6

CTATACAAAATGGCACAGCTTGAGCAAAAAGATGGCGATATTT

F3

AATCTGCAGCCGTTCTTTTGCAAT

3–1

GAACGGCTGCAGATTTATCTGCCGCAGTTGCGTAAAGATATTCT

3–2

GAACGGCTGCAGATTGTACAAATTGATCCTGAGATGGTAACCGTACA

In order to construct a series of egfp fusions carrying C-terminal, N-terminal, or middle part truncated MinE, 20 primers were designed. Oligos 1–1 to 1–6 as the forward primers and F1 as the reverse (Table 1) were used to construct vectors pMinE1–78, pMinE1–59, pMinE1–45, pMinE1–34, pMinE1–25, and pMinE1–16 (Fig. 1), in which the reverse primer shares 15 homolog nucleotides in the 5′ end with the forwards. The rPCR was performed using the plasmid pGM as template, and then the amplicons were self-ligated using In-Fusion® Dry-Down PCR Cloning Kit to generate a series of truncated MinE from C-terminal. The similar method was applied to construct the eGFP fused with N-terminal truncated or middle-region-deleted MinE (Fig. 2), pMinE11–88, pMinE18–88, pMinE22–88, pMinE40–88, pMinE55–88, and pMinE64–88 using the primers 2–1 to 2–9 as the forward and F2 as the reverse, or pMinE1–24/38–88 and pMinE1–24/54–88 with the primers 3–1 and 3–2 as the forwards and F3 as the reverse. All the vectors were used to transform E. coli DH5α cells.
Fig. 1

Schematic representation of oligonucleotide binding sites for deleting C-terminal fragments from MinE on the plasmid pGM

Fig. 2

Diagram for the construction of egfp-minE fusions. a Secondary structural elements and protein domains in MinE. b Schematic representation of the position of deleted nucleotides in egfp-minE fusions. The numbers in superscripts of MinE refer to the deleted amino acid residues

DNA sequencing

All the constructs were sequenced with the proper primers to verify that no mutations had been introduced during the construction. The process is similar as Fu et al. mentioned (Fu et al. 2014). The BigDye_ Terminator v3.1 Cycle-Sequencing Kit and ABI 3130 automated sequencer were used for sequencing.

Epifluorescence microscope

The colonies of transformed E. coli were cultured on LB medium supplemented with ampicillin (100 μg/ml) overnight, and then used to visualize the cell phenotypes with ×100 objective, 540–550-nm excitation filter and 590-nm barrier filter for observation under an epifluorescence microscope (Olympus BX50). The pictures were taken by a Nikon Ltd camera and the Image Pro Plus 5.0 software for treatment.

Results and discussion

System construction for protein localization test

Firstly, we established a system in which the recombinant protein eGFPp-MinE showed similar functions and similar subcellular localization as the chromosomal MinE (chrMinE) while the expression of eGFP-MinE does not apparently affect cell shape. The plasmid pBluscriptSK(−) was used for modification, which is a multiple-copy plasmid with inducible strong promoter. Thus, a constitutive promoter Pfla from Helicobacter pylori has been introduced into this plasmid to replace the Plac or much closer to the ORF of recombinant proteins. Then, the plasmids that contain the complete MinE gene sequence in-fusion to egfp as the minE-egfp or egfp-minE under both promoters Plac and Pfla (Pfla much closer to the ORF than Plac) resulted in the plasmids pMG or pGM, respectively, or the plasmid containing egfp-minE under a unique promoter Pfla (by replacing the Plac promoter in pBluescript) was called pFGM. E. coli DH5alpha was transformed with pMG, pGM, and pFGM, respectively. All three cases displayed the same MinE phenotype (Fig. 3) (Hale et al. 2001; Fu et al. 2001), a fluorescence signal at the pole from the cells, indicating that the recombinant proteins egfp-minE or minE-egfp under both promoters or only Pfla showed similar function and the same feature of subcellular localization as MinE. Thus, this system can be used for protein localization test. On the other hand, as there is no difference in the phenotypes between pGM and pMG, pGM was used as the base for the following recombinant truncated MinE construction.
Fig. 3

eGFP, eGFP-MinE, or MinE-eGFP localization feature in E. coli. a pBS80, b pMG, c pGM, and d pFGM

The phenotypes related to MinE in E. coli have been documented, such as gene deletion and overexpression. If E. coli cells lack MinE protein, the MinCD inhibitory complex is placed throughout the cell inhibiting the Z-ring formation in the whole cell (Hu and Lutkenhaus 1999). Hence, the cell division is inhibited (de Boer et al. 1989; Fu et al. 2001), resulting in non-septate filamentous cells (Raskin and de Boer 1999). MinE overexpression leads to chromosome-less minicell formation, similar to the cells lacking the three Min genes (Min¯ phenotype) (de Boer et al. 1989; Labie et al. 1990). In our study, the chrMinE gene was not deleted, whereby transformed bacteria hold basic expression levels of MinE in addition to the recombinant MinE expressed from plasmids. Thus, the results here reflect the effect of egfp-minE expression (whole or truncated MinE) in competition with the inherent MinE for the targets in a cell.

The MinE overexpression in E. coli counteracts the inhibitory action from MinCD complex, causing septation either in the pole or middle cell and leading to the production of chromosome-less minicells (de Boer et al. 1989; King et al. 1999; Pichoff et al. 1995; Labie et al. 1990; Ghasriani et al. 2010). This was consistent with the E. coli cells transformed with pFGM, pMG, and pGM (all of them expressed the whole MinE gene), which have shown some minicells, indicating that the fusion proteins (N- and C-terminal fusions) in this study have the similar function as chromosome-encoded MinE and do not affect its subcellular localization but still overexpressed.

Protein eGFP fused with MinE, truncated from C-terminal and phenotypes of fusion protein localization

The series of eGFP fused with incomplete MinE fusion proteins, partially deleted from C-terminal, was constructed in the same vector (pGM), as shown in Fig. 2. The localization feature of fluorescent MinE can be observed in the transformants with the plasmids containing partial deletions of MinE under fluorescent microscopy.

The fusion proteins from MinE1–78 (the number on superscript indicates the presence of MinE amino acids in fusion protein) to MinE1–16 lack the C-terminal residues, from deletion of 10 to 72 amino acids except the stop codon. The recombinant protein signals in cells of transformants are similar to those of transformants with pMG, pGM, and pFGM (Fig. 4a–d). Although the transformants with pMinE1–25 pMinE1–16 still showed the localization feature as transformants with pMG or pGM, some fusion protein diffused throughout the cells is evident (Fig. 4e, f).
Fig. 4

Bacteria transformed with plasmids containing C-terminal truncated minE. a MinE1–78, b MinE1–59, c MinE1–45, d MinE1–34, e MinE1–25, and f MinE1–16

These fusion proteins in the plasmids from pMinE1–78 to pMinE1–34 partially or completely lack the TSD but the anti-MinCD domain is still present. Previous studies have shown that MinE1–34 and MinE1–53 protein can counteract the MinCD inhibitory action in the MinE mutants (Pichoff et al. 1995; Rowland et al. 2000). Thus, we understand that if the C-terminal truncated MinE has the ability for dimerization (homodimers or heterodimers), these dimers can perform the anti-MinCD as homodimers of chromosomal MinE; otherwise, they cannot compete with chromosomal MinE dimers. MinE1–25 and MinE1–16 lack partially the anti-MinCD domain, particularly the β1 secondary element (Shih and Zheng 2013; Park et al. 2011), which is relevant to the binding ability to the MinD (Park et al. 2011), in addition to the absence of TSD, and have lost the ability for heterodimer formation with chrMinE and their anti-MinCD function. However, the protein localization was affected to a degree due to some florescent signal distributed all over the cells when transformed with pMinE1–25 and pMinE1–16, implying that their structures could be unstable.

Protein eGFP fused with MinE, truncated from N-terminal and phenotypes of fusion protein localization

The series of eGFP fusion proteins from “MinE11–88” to “MinE64–88” lacks the N-terminal residues of MinE (Fig. 2). Bacteria transformed with vectors expressing fusion proteins from “MinE11–88” to “MinE22–88” showed elongated filaments and clear but distinct localization feature from the MinEs truncated from C-terminal (Fig. 5a–c). These expressed proteins lack the complete membrane-binding domain. “MinE32–88” lacks both the membrane-binding and the anti-MinCD domain and some fluorescence signal distributed throughout the cell whereas some stronger signal was localized close to the pole or in some other regions of filamentous cells, however, in much smaller cell size (Fig. 5d). Bacteria expressing MinE38–88 showed the filaments with the similar cell size to the strain expressed with “MinE32–88,” and fluorescent signal was localized in the poles and some other specific sites of cells (Fig. 5e). The cells expressing MinE40–88 showed the signal feature as those with “MinE11–88,” “MinE18–88,” and “MinE22–88” but with more diffused recombinant proteins (Fig. 5f). The phenotype of cells with MinE48–88 (Fig. 5g) was similar to the bacteria expressing MinE38–88. The cells with the expression of MinE55–88 and MinE64–88 yielded fluorescent signal that almost lost the localization and was distributed throughout the cell (Fig. 5h, i, respectively). Actually, a small part of those fusions still showed similar localization close to the pole as MinE. The size of cells with MinE55–88 was similar to the cells expressing N-terminal truncated MinE fusions “MinE11–88,” “MinE18–88,” and “MinE22–88” aforementioned and the size of cells with MinE64–88 showed much smaller.
Fig. 5

Bacteria transformed with plasmids containing N-terminal truncated minE. a MinE11–88, b MinE18–88, c MinE22–88, d MinE32–88, e MinE38–88, f MinE40–88, g MinE48–88, h MinE55–88, and i MinE64–88

The fusion protein from MinE11–88 contains the entire TSD without the membrane-binding domain. It should have the ability to form the dimers, homodimers, or heterodimers with inherent MinE (King et al. 2000; Ghasriani and Goto 2011). Considering that the cells are long, non-separate filaments, it can be inferred that the incomplete MinEs are able to form heterodimers with the inherent complete MinE or to form its homodimers with no complete anti-MinCD function. It is reasonable that if these dimers involved in incomplete MinE have complete anti-MinCD function, the cell phenotype should be the same case of overexpression of MinE or the expression of some C-terminal-truncated MinEs in the present study (Fig. 4a–d). This result also confirmed that the membrane-binding domain is the precondition for anti-MinCD function of MinE. In MinE dimers, one membrane-binding domain binds to the cell membrane that allows the homodimer to move in a similar way to Tarzan swinging on vines (Park et al. 2011), reaching a nearby membranal MinCD complex (Hu et al. 2003), where the ATPase activity leads to the release of membranal MinCD complex (Bramkamp and van Baarle 2009; Laloux and Jacobs-Wagner 2014). Therefore, a heterodimer formed by one chrMinE and one truncated MinE lacking the membrane-binding domain is unable to perform this function, decreasing the ability of cells to inhibit the MinCD effect, leading to filamentation of cells and a localization feature with fluorescence in a zebra pattern (multiple rings of fusion protein) (Fu et al. 2001; Raskin and de Boer 1997). The fusions MinE18–88 and MinE22–88, which contain the incomplete anti-MinCD domain (King et al. 1999) and the complete TSD, still have the ability to form heterodimers with chrMinE because they showed the same phenotype. The fusions MinE32–88 and MinE38–88, only containing the complete TSD region, resulted in the short filamentous cells and minicells, similar to the phenotype of MinE36–88 that was confirmed not to be able to form chrMinE/MinE36–88 hetero-oligomers (Zhang et al. 1998), but the homodimers of MinE32–88 and MinE38–88 still have the weak competition with chrMinE for the topological specificity of septal placement. The fusion MinE40–88 showed the same phenotype as MinE18–88 and MinE22–88, suggesting that MinE40–88 can also form chrMinE/MinE40–88 hetero-oligomers. MinE48–88, containing the most part of TSD region, leads to the similar cell phenotype as MinE32–88 and MinE38–88, and they could share the similar conformation. King et al. mentioned that the mutations Y38F and D45A in MinE made the MinE to lose topological specificity (King et al. 2000). These results indicate that TSD region (32–48) responsible for MinE dimerization is sensitive for conformation change. However, how do the fusions MinE32–88, MinE38–88, and MinE48–88 affect the initiation site for cell division different from other N-terminal truncated MinE fusions? It is still a question to be clarified in the future.

The transformants carrying the plasmids that expressed the truncated proteins MinE55–88and MinE64–88 showed that florescent signal occurred throughout the cell and the cell was still in the filamentous shape, implying that most of the fusions lost the localization ability and the other fusions localized still interfere the function of chrMinE. For example, the heterodimers with chrMinE can compete with the homodimers from chrMinE due to the presence of one complete β3 domain and one complete or partial β2 regions in the truncated MinEs. It also suggests that these structures are unstable. As MinE is more truncated in this region, much less fusions have the ability to interfere chrMinE; the cell is much closer to the normal shape, and more eGFP fusion molecules do not localize and scatter throughout the cell.

Protein eGFP fused with MinE, truncated in the middle of gene and phenotypes of fusion protein localization

Bacteria transformed with pMinE1–24/38–88 (Fig. 6a) still displayed filamentous phenotypes and fluoresces in a zebra pattern (similar to Fig. 5a), and cells transformed with pMinE1–24/54–88 displayed principally similar localization to those transformed with MinE1–25 and MinE1–16 but without florescence signal diffusion (Fig. 6b). Both proteins lack the partial anti-MinCD TSD domain.
Fig. 6

Bacteria transformed with plasmids containing the middle region-truncated minE. a MinE1–24/38–88 and b MinE1–24/54–88

Comparing the phenotype of the fusion MinE1–24/38–88 with those of MinE1–25 and Min38–88, the phenotype of the fusion MinE1–24/38–88 is similar neither to MinE1–25 nor Min38–88 but to MinE11–88, MinE18–88, and MinE22–88. Thus, it means that the fusion MinE1–24/38–88 keeps the conformation as MinE11–88and has the ability to form dimers (homodimer of MinE1–24/38–88 or heterodimer with chrMinE) but with incomplete anti-MinCD domain, which could compete with chrMinE homodimers, resulting in the filamentous phenotype. However, the fusion MinE1–24/54–88 showed completely different phenotype from Min55–88 but similar to minE1–25 without signal protein diffusion, suggesting that its localization principally depends on the region of 1–25 amino acids, not 55–88 amino acids. Considering that a lot of fusion molecules of MinE55–88 were scattered distribution throughout the cells (Fig. 5), it can be concluded that the region of 55–88 amino acids is not essentially involved in MinE subcellular localization although the structure of some fusions is unstable and still involved in ChrMinE competition or other unknown process of anti-MinCD. In other words, three functional domains in the region of 1–54 amino acids principally decide the MinE positioning in cells.

Thus, the essence of protein subcellular localization is the consequences of a protein interacted with other biological macromolecules in cells. The present method gave a model for essential gene study. If the protein has several factors involved in the localization, the localization signal disappears as all the factors are eliminated. When the protein forms polymers, it can be considered as an independent factor if the polymers are related to protein localization. If it is not the house-keeping gene, the series of truncated genes fused with egfp can be expressed in the gene-chromosomal deletion mutant and show the localization signal as one of localization factors is expressed. Thus, each strategy to study the sequences that determine the protein localization has its own advantage, using either of them or both strategies that depend on the complexity of localization factors of the gene.

In summary, the localization and function of a protein are strongly related but distinct. The adequate localization is the base for a protein to work in cells. The function is always the focus for protein research whereas the subcellular localization study can also provide new information for the protein. If a protein has a specific subcellular location, the sequence responsible for it should exist in the protein. In the present study, the sequences related to the subcellular localization of MinE in cells were determined (1–54 amino acids), which contains the functional regions as membrane-binding domain, anti-MinCD domain, and some of dimerization domain. Without the dimerization domain, the truncated MinE almost cannot interfere the function of chromosomal MinE. After the membrane-binding and anti-MinCD regions in N-terminal truncated MinEs were eliminated, the rest MinE fragments have different ability to form heterodimers between chromosomal MinE and truncated MinE, derived from the phenotypes of cells. In some cases, the structures of truncated proteins are unstable with fluorescent signals distributed throughout the cells. These results imply that protein subcellular localization is complicated for MinE, and its mechanism in detail is still unknown. For example, what is the biological significance of MinE conformation change so sensitive? What are the conditions that could lead to the conformation change? How do the homodimers of MinE32–88 and MinE38–88 or MinE48–88 realize to interfere chrMinE for the topological specificity of septal placement? What molecule does the MinE membrane domain attach? Thus, the studies on subcellular localization can find some new characteristics of a protein, which helps us to understand better the protein activity in the cells. In general, the series of N-terminal and C-terminal deletion and some intermediate deletion of a protein can provide adequate information on the sequences for protein localization. This method can be generally used for protein localization signal sequence analysis, especially for essential protein analysis.

Notes

Funding information

This work was supported by the Consejo Nacional de Ciencia y Tecnología (CONACyT)-México (Grant No. 168541) and Secretaría de Investigación y Posgrado del Instituto Politécnico Nacional, México (No. 20151373). Miguel Ángel Pérez-Rodríguez held scholarships from CONACyT. X. Guo and Mario A. Rodriguez Perez hold scholarships from COFAA-IPN.

Compliance with ethical standards

Competing interests

The authors declare that they have no conflict of interest.

Supplementary material

10123_2018_1_MOESM1_ESM.png (95 kb)
Supplementary Figure 1 (PNG 95.3 kb)

References

  1. Bramkamp M, van Baarle S (2009) Division site selection in rod-shaped bacteria. Curr Opin Microbiol 12:683–688CrossRefPubMedGoogle Scholar
  2. Cormack BP, Valdivia RH, Falkow S (1996) FACS-optimized mutants of the green fluorescent protein (EGFP). Gene 173:33–38CrossRefPubMedGoogle Scholar
  3. de Boer PA, Crossley RE, Rothfield LI (1989) A division inhibitor and a topological specificity factor coded for by the minicell locus determine proper placement of the division septum in E. coli. Cell 56:641–649CrossRefPubMedGoogle Scholar
  4. Emanuelsson O, Nielsen H, Brunak S, Von Heijne G (2000) Predicting subcellular localization of proteins based on their N-terminal amino acid sequence. J Mol Biol 300:1005–1016CrossRefPubMedGoogle Scholar
  5. Fu X, Shih Y-L, Zhang Y, Rothfield LI (2001) The MinE ring required for proper placement of the division site is a mobile structure that changes its cellular location during the Escherichia coli division cycle. Proc Natl Acad Sci U S A 98:980–985CrossRefPubMedPubMedCentralGoogle Scholar
  6. Fu Y, Zepeda-Gurrola RC, Aguilar-Gutiérrez GR, Lara-Ramírez EE, De Luna-Santillana EJ, Rodríguez-Luna IC, Sánchez-Varela A, Carreño-López R et al (2014) The detection of inherent homologous recombination between repeat sequences in H. pylori 26695 by the PCR-based method. Curr Microbiol 68:211–219CrossRefPubMedGoogle Scholar
  7. Ghasriani H, Goto NK (2011) Regulation of symmetric bacterial cell division by MinE: what is the role of conformational dynamics? Commun Integr Biol 4:101–103CrossRefPubMedPubMedCentralGoogle Scholar
  8. Ghasriani H, Ducat T, Hart CT, Hafizi F, Chang N, Al-Baldawi A, Ayed SH, Lundström P et al (2010) Appropriation of the MinD protein-interaction motif by the dimeric interface of the bacterial cell division regulator MinE. Proc Natl Acad Sci U S A 107:18416–18421CrossRefPubMedPubMedCentralGoogle Scholar
  9. Hale CA, Meinhardt H, de Boer PA (2001) Dynamic localization cycle of the cell division regulator MinE in Escherichia coli. EMBO J 20:1563–1572CrossRefPubMedPubMedCentralGoogle Scholar
  10. Hsieh CW, Lin TY, Lai HM, Lin CC, Hsieh TS, Shih YL (2010) Direct MinE–membrane interaction contributes to the proper localization of MinDE in E. coli. Mol Microbiol 75:499–512CrossRefPubMedGoogle Scholar
  11. Hu Z, Lutkenhaus J (1999) Topological regulation of cell division in Escherichia coliinvolves rapid pole to pole oscillation of the division inhibitor MinC under the control of MinD and MinE. Mol Microbiol 34:82–90CrossRefPubMedGoogle Scholar
  12. Hu Z, Lutkenhaus J (2000) Analysis of MinC reveals two independent domains involved in interaction with MinD and FtsZ. J Bacteriol 182:3965–3971CrossRefPubMedPubMedCentralGoogle Scholar
  13. Hu Z, Saez C, Lutkenhaus J (2003) Recruitment of MinC, an inhibitor of Z-ring formation, to the membrane in Escherichia coli: role of MinD and MinE. J Bacteriol 185:196–203CrossRefPubMedPubMedCentralGoogle Scholar
  14. King GF, Rowland SL, Pan B, Mackay JP, Mullen GP, Rothfield LI (1999) The dimerization and topological specificity functions of MinE reside in a structurally autonomous C-terminal domain. Mol Microbiol 31:1161–1169CrossRefPubMedGoogle Scholar
  15. King GF, Shih Y-L, Maciejewski MW, Bains NP, Pan B, Rowland SL, Mullen GP, Rothfield LI (2000) Structural basis for the topological specificity function of MinE. Nat Struct Mol Biol 7:1013–1017CrossRefGoogle Scholar
  16. Labie C, Bouché F, Bouché J (1990) Minicell-forming mutants of Escherichia coli: suppression of both DicB-and MinD-dependent division inhibition by inactivation of the minC gene product. J Bacteriol 172:5852–5855CrossRefPubMedPubMedCentralGoogle Scholar
  17. Laloux G, Jacobs-Wagner C (2014) How do bacteria localize proteins to the cell pole? J Cell Sci 127:11–19CrossRefPubMedPubMedCentralGoogle Scholar
  18. Park K-T, Wu W, Battaile KP, Lovell S, Holyoak T, Lutkenhaus J (2011) The Min oscillator uses MinD-dependent conformational changes in MinE to spatially regulate cytokinesis. Cell 146:396–407CrossRefPubMedPubMedCentralGoogle Scholar
  19. Pazos M, Casanova M, Palacios P, Margolin W, Natale P, Vicente M (2014) FtsZ placement in nucleoid-free bacteria. PLoS One 9:e91984CrossRefPubMedPubMedCentralGoogle Scholar
  20. Pichoff S, Vollrath B, Touriol C, Bouché JP (1995) Deletion analysis of gene minE which encodes the topological specificity factor of cell division in Escherichia coli. Mol Microbiol 18:321–329CrossRefPubMedGoogle Scholar
  21. Raskin DM, de Boer PA (1997) The MinE ring: an FtsZ-independent cell structure required for selection of the correct division site in E. coli. Cell 91:685–694CrossRefPubMedGoogle Scholar
  22. Raskin DM, de Boer PA (1999) Rapid pole-to-pole oscillation of a protein required for directing division to the middle of Escherichia coli. Proc Natl Acad Sci U S A 96:4971–4976CrossRefPubMedPubMedCentralGoogle Scholar
  23. Rowland S, Fu X, Sayed M, Zhang Y, Cook W, Rothfield L (2000) Membrane redistribution of the Escherichia coli MinD protein induced by MinE. J Bacteriol 182:613–619CrossRefPubMedPubMedCentralGoogle Scholar
  24. Rudner DZ, Losick R (2010) Protein subcellular localization in bacteria. Cold Spring Harb Perspect Biol 2:a000307CrossRefPubMedPubMedCentralGoogle Scholar
  25. Shapiro L, McAdams HH, Losick R (2009) Why and how bacteria localize proteins. Science 326:1225–1228CrossRefPubMedGoogle Scholar
  26. Shih YL, Zheng M (2013) Spatial control of the cell division site by the Min system in Escherichia coli. Environ Microbiol 15:3229–3239CrossRefPubMedGoogle Scholar
  27. Zhang Y, Rowland S, King G, Braswell E, Rothfield L (1998) The relationship between hetero-oligomer formation and function of the topological specificity domain of the Escherichia coli MinE protein. Mol Microbiol 30:265–273CrossRefPubMedGoogle Scholar
  28. Zhao C-R, De Boer P, Rothfield LI (1995) Proper placement of the Escherichia coli division site requires two functions that are associated with different domains of the MinE protein. Proc Natl Acad Sci U S A 92:4313–4317CrossRefPubMedPubMedCentralGoogle Scholar
  29. Zhou H, Schulze R, Cox S, Saez C, Hu Z, Lutkenhaus J (2005) Analysis of MinD mutations reveals residues required for MinE stimulation of the MinD ATPase and residues required for MinC interaction. J Bacteriol 187:629–638CrossRefPubMedPubMedCentralGoogle Scholar

Copyright information

© Springer International Publishing AG, part of Springer Nature 2018

Authors and Affiliations

  • Miguel Á. Pérez-Rodríguez
    • 1
    • 2
  • Isabel Cristina Rodríguez-Luna
    • 2
  • Ricardo Carreño-López
    • 3
  • Edgar E. Lara-Ramírez
    • 4
  • Mario A. Rodríguez-Pérez
    • 2
  • Xianwu Guo
    • 2
  1. 1.Departamento de BotánicaUniversidad Autónoma Agraria Antonio NarroSaltilloMexico
  2. 2.Centro de Biotecnología GenómicaInstituto Politécnico NacionalCd. ReynosaMexico
  3. 3.Centro de Investigaciones en Ciencias MicrobiológicasBenemérita Universidad Autónoma de PueblaPueblaMexico
  4. 4.Unidad de Investigación Biomédica de ZacatecasInstituto Mexicano del Seguro Social (IMSS)ZacatecasMexico

Personalised recommendations