*2.3.2 Secondary sources of data*

The secondary sources of data were AC and DC minutes in the college and the departments, and the document from Quality Assurance Directorate Office on the teaching-learning performance in the college.

*Teachers' Ethical Professional Practices in Higher Education Institutions: An… DOI: http://dx.doi.org/10.5772/intechopen.109651*

### **2.4 Sample and sampling technique**

In this study, six students were selected for the focus group discussion, and four teachers and four department heads were selected for the in-depth interview purposively. Lodging complaints to the department was used as criteria for selecting the students. Department heads were included into the study because they were the ones who handle complaints against teachers brought to their respective departments, and members of Academic Council were included because complaints were seen for further deliberation and decisions at this stage. In this study, purposive sampling technique was employed for the reason that this sampling design enables the researchers to get information-rich study participants (**Table 1**) [23].

#### **Table 1.**

*Sample and sampling technique.*

#### **2.5 Instruments of data collection**

In-depth interview, focus group discussion, and documents analysis were used to collect data. An in-depth interview was employed because it enables to explore indepth a respondent's point of view, experiences, feelings, and detailed personal information [24]. Document analysis was carried out because it was used as a means of triangulation to provide a confluence of evidence that breeds credibility [25]. Focus group discussion was employed because it helps to probe deeper into certain things have or not have occurred, and to explore un anticipated issues [26].

#### **2.6 Method of data analysis**

The analysis of data was carried out as follows. First, transcriptions of data collected through in-depth interview and focus group discussion were read and re-read several times. Then, data were coded and categorized into their similarities, and eventually, themes were formed, and finally, thick description of the data was done.


**Mutation of** *Inositol 1,3,4-trisphosphate 5*/*6-kinase6* **Impairs Plant Growth and Phytic Acid Synthesis in Rice**

### **Meng Jiang 1,2,3 , Yang Liu 1, Yanhua Liu 1, Yuanyuan Tan 1, Jianzhong Huang 1,3 and Qingyao Shu 1,2,\***

Received: 20 March 2019; Accepted: 24 April 2019; Published: 29 April 2019

**Abstract:** Inositol 1,3,4-trisphosphate 5/6-kinase (ITPK) is encoded by six genes in rice (*OsITPK1-6*). A previous study had shown that nucleotide substitutions of *OsITPK6* could significantly lower the phytic acid content in rice grains. In the present study, the possibility of establishing a genome editing-based method for breeding low-phytic acid cultivars in rice was explored, in conjunction with the functional determination of OsITPK6. Four *OsITPK6* mutant lines were generated by targeted mutagenesis of the gene's first exon using the CRISPR/Cas9 method, one (*ositpk6\_1*) with a 6-bp in-frame deletion, and other three with frameshift mutations (*ositpk6\_2*, \_*3*, and \_*4*). The frameshift mutations severely impaired plant growth and reproduction, while the effect of *ositpk6\_1* was relatively limited. The mutant lines *ositpk6\_1* and \_*2* had significantly lower levels (−10.1% and −32.1%) of phytic acid and higher levels (4.12- and 5.18-fold) of inorganic phosphorus compared with the wild-type (WT) line. The line *ositpk6\_1* also showed less tolerance to osmotic stresses. Our research demonstrates that mutations of *OsITPK6*, while effectively reducing phytic acid biosynthesis in rice grain, could significantly impair plant growth and reproduction.

**Keywords:** genome editing; growth; *ositpk6*; phytic acid; rice


**1. Introduction: historical perspective**

The intrinsic awareness in structure-properties relationship of solids was firstly introduced on the study of minerals and inorganic compounds and has been preserved over time [1]. Within this frame, the first step on the polymorphism phenomenon was unknowingly given by Klaproth [2] in 1788 identifying three different crystalline phases of calcium carbonate (calcite, vaterite and aragonite). In 1819, Mitscherlich commenced his research on phosphates and arsenates of potassium (KH2PO4 and KH2AsO4) and established their complete morphological similarity, noting that *they crystallize in similar forms*. His subsequent results with the corresponding potassium salts (NH4H2PO4 and NH4H2AsO4) confirmed that observation and led him to pose that *there do exist bodies of dissimilar chemical composition having the same crystalline form*. He not only recovered the work of Whollaston with orthorhombic carbonates and sulfates of barium, strontium and lead, who already noticed this phenomenon in 1812 but also, extended to rhombohedral carbonates of calcium, magnesium, iron and manganese, and to sulfates of iron, copper, zinc, magnesium, nickel and cobalt. Curiously, he also evinced the basis of *seeding* at realizing that *having two substances both able to crystallize in various forms, the presence of one during the crystallization of the other will force the crystallization of the latter in the same form*. Then, he moved to Stockholm with Berzelius with whom he delved deeply into phosphates and arsenates and forged the concept of

*isomorphism*. Last but not least, he reported two distinct crystallized forms of sulfur in 1826 demonstrating *a clear case of an element which could be made to crystallize in two different systems of symmetry at will, by merely changing the crystallization conditions.* Therefore, Mitscherlich was identified as the one who took the first step towards the rise of *polymorphism* [3]. Shortly after, in 1824, the contradictory results garnered by Liebig [4] and Wöhler [5] during their research on silver fulminate and silver cyanate triggered the conflict which led them to be colleagues and to pose the following dilemma: *can two compounds with the same composition have different physical properties?*. Their results evinced two Ag salts with the same composition but different physical properties, which did not go unnoticed by Wöhler's master Berzelius, who merged these results with those of Mitscherlich. It is unclear who was the first to conceive the notion of *polymorphism* but this crucial period of 1820–1832 was mainly drawn by Mitscherlich [6–8] who shed light on this phenomenon, even though the concept was still vague. It was during these years when Berzelius [9] proposed the concept of *isomerism* (1831–1832) and it took until 1832 for Wöhler and Liebig [10] to report the first case of *polymorphism* in an organic compound. The awareness of *isomerism* set the beginning of structural chemistry, broadening the knowledge and understanding of organic structures.

Since observation was the essential tool to identify *polymorphism*, this research drastically changed with the accessibility and wide spreading use of the microscope, but it was not until 1839 when Frankenheim [11] introduced the first principles defining *polymorphism* and *a postriori*, Mallard [12] set the structural basis of *polymorphism* in 1876, relating differences in physical properties with different arrangements. One of the most remarkable contribution during this period was the "Rule of steps" or "Law of successive reactions" from Ostwald [13] in 1897. He pointed that during a succession of polymorphic forms, those appeared later are generally more stable. Despite not being considered a rule, it is still valid as a general observation. But the two major queries raised by Buerger and Bloom [14] in 1937 were still unanswered: what causes the formation of different phases of a substance and which factors determine them?.

The narrow link between crystallography and *polymorphism* was forged by Tamman [15] in 1926 and settled with the first polymorphic X-ray crystal structure determination of an organic compound, resorcinol, published by Robertson and Ubbelohde [16] in 1939. Despite this achievement, the next decades passed without a better understanding of *polymorphism,* being underrated until 1965 when McCrone [17] conducted a comprehensive study in which he defined a polymorph as: "*a solid crystalline phase of a given compound resulting from the possibility of at least two different arrangements of the molecules of that compound in the solid state*" and published a review in 1969 about the importance of such phenomenon in the pharmaceutical outlook [18]. Since the introduction of the term *allotrope* by Berzelius in 1841, *Polymorphism* had been taken with *allotropy* on the same meaning. But, it was not until the 1990s, when Sharma [19] and Reinke [20] set the differences between them: *polymorphism* occurs in chemical compounds while *allotropy* occurs in chemical elements. This work was crowned *a posteriori* by Dunitz's [21, 22] crystal description contribution.

In the 1970s, the works of Schmidt [23] and Paul and Curtin [24, 25] grounded the flourish of solid-state chemistry and precede the breakthrough of conceiving polymorphism. They served as inspiration to Bernstein and Desiraju, who laid the foundation for recent supramolecular chemistry. In 1978, Bernstein [26] changed the landscape of *polymorphism* by rationalizing the study of crystal packing forces on molecular conformations of polymorphs and later in 1990, together with Etter [27–30], applied their graph set descriptors and provide guidelines to understand polymorphic transformations [31]. Bernstein compiled most of this historical

#### *Polymorphism and Supramolecular Isomerism: The Impasse of Coordination Polymers DOI: http://dx.doi.org/10.5772/intechopen.96930*

results in his book "*Polymorphism in Molecular Crystals*", which the authors encourage reading [32]. Likewise, Desiraju achieved substantial progress in this field during his studies of structure-properties relationship of organic solids mainly of pharmaceutical interest, emphasizing and aiming its importance in this industry as reflected in his book "*Crystal Engineering: A Textbook*" [33]. Both channeled their polymorphism vision in terms of supramolecular chemistry.

During 1990s, computational chemistry went hand in hand, achieving methodologies capable to reproduce experimental results and enabling even crystal structure predictions. The first attainment was obtained by using Williams' software [34] that met the main handicap hitherto, the identification of lattice energy minimums. Subsequent years, many computational approaches were developed facing with computer-generated structures for prediction, to the extent that in 1999 a collaborative workshop held at the Cambridge Crystallographic Data Centre (CCDC) [35] brought together the benchmark computational groups of this period to provide an objective assessment of the possibilities of crystal structure prediction. The results gathered in this event were clearly summarized *a posteriori* in a paper published by Lommerse [36]. Further advances on prediction methods accuracy as well as the implementation of Density Functional Theory (*DFT*) and Machine Learning (*ML*) can be found in Spark's review [37].

Despite the basis of polymorphism were already defined at the end of the twentieth century, the increasing advances on X-ray diffraction techniques and crystallization methods afforded the determination and analysis of metal–organic structures, especially *coordination polymers*. *Polymorphism* of metal–organic complexes was still unexplored and these new class of materials, from which polymorphic structures grew exponentially, required classification and awareness. This impasse was encouraged by renowned researchers as Sharma [38], Ciani [39], Rogers [40] and Zaworotko [41]. In this regard, the same Zaworotko was who published a review in 2001 emphasizing the difference between *polymorphism* and *supramolecular isomerism* and underlined the link between them in organic and metal–organic networks [42].


**Quinolone Resistance in Tuberculosis Treatment: A Structural Overview**

Claudine Mayer1,2,3 and Alexandra Aubry4,5,6 *1Département de Biologie Structurale et Chimie, Institut Pasteur, Paris 2URA 2185 du CNRS, Paris 3Univ Paris Diderot, Sorbonne Paris Cité (Cellule Pasteur), Paris 4UPMC Univ Paris 06, ER5, Laboratoire de Bactériologie, Paris 5AP-HP, Hôpital Pitié-Salpêtrière, Laboratoire de Bactériologie-Hygiène, Paris*

*6CNR des Mycobactéries et de la Résistance des Mycobactéries aux Antituberculeux, Paris France*

## **1. Introduction**

290 Understanding Tuberculosis – New Approaches to Fighting Against Drug Resistance

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#### **1.1 Fluoroquinolones in tuberculosis treatment**

Fluoroquinolones are key antibiotics for the second-line treatment of multidrug-resistant tuberculosis (MDR-TB), i.e. resistant to the two most powerful antituberculous drugs currently available, isoniazid and rifampicin (WHO, 2001; 2011). Unfortunately, extensively drug resistant strains (XDR-TB), defined as MDR-TB with bacillary resistant to any fluoroquinolones and to one of the three injectable second line anti-TB drugs (amikacin, kanamycin or capreomycin), are emerging worldwide (Center for Disease Control and Prevention, 2006). XDR-TB strains constitute a major concern for public health since they are virtually untreatable and responsible for up to 100% of deaths. Besides being increasingly popular in treatment of tuberculosis complicated by intolerance or by relative contraindication for first-line drugs, fluoroquinolones are under studies to shorten the duration of treatment or replace first line drugs in susceptible tuberculosis. Newer fluoroquinolones such as moxifloxacin have already demonstrated potential for shortening treatment duration (Rustomjee et al., 2008; Conde et al., 2009; Dorman et al., 2009).

Fluoroquinolones are also one of the most widely prescribed antibiotics as they are generally well tolerated with high oral bioavailability plus broad-spectrum antibacterial activities against genitourinary infections and against common respiratory tract pathogens. Structurally, these drugs contain a quinoline ring system and hence are given the name quinolones (Fig. 1). After the first generation of drugs were found to be active, it was noted that a fluorine atom at the 6-position of the quinoline ring imparted greater potency, and hence the second generation of drugs was called the fluoroquinolones (Fig. 1).

Quinolone Resistance in Tuberculosis Treatment: A Structural Overview 293

5). The BRD contains the catalytic tyrosine responsible for the cleavage and religation of the

Fig. 2. Schematic representation of the sequence and domain organization of type II topoisomerases found in eukaryotes, bacteria, virus and archaea. The type IIA

reunion domain (BRD) of type IIA topoisomerase.

open clamp form (Fig. 3).

topoisomerase family includes bacterial DNA gyrase and topoisomerase IV (Topo IV), eucaryal and viral topoisomerases (Topo II), whereas the type IIB topoisomerase family includes archaeal topoisomerase VI (Topo VI), and their homologues in plants and a few protists and bacteria. Topo IIA and Topo IIB are both formed by the association of two subunits A and B. Bacterial type IIA and type IIB topoisomerases are A2B2 heterotetramers, whereas the eukaryotic Topo II is a heterodimer, with the A and B subunits fused in a single polypeptide (Nter-B-A-Cter). The names of the four conserved domains are indicated. The winged helix domain (WHD) in the type IIB topoisomerase corresponds to the breakage-

Crystallographic studies of individual domains show that the structure of the BRD is a heart-shaped arrangement with two dimer interfaces (Morais-Cabral et al., 1997), that the CTD displays a spiral six-bladed β-pinwheel structure (Corbett et al., 2004) and that the ATPase domain belongs to the GHKL ATPases, a broad family of enzymes with a common fold unrelated to other canonical ATP-binding folds (Brino et al., 2000) (Fig. 3). The Toprim domain interacts with the A subunit, possesses a magnesium-binding site and is essential for DNA binding (Fig. 3). Although no structure for a fully intact active type IIA topoisomerase has been determined yet, combination of structural and biochemical studies of the individual domains has lead several authors to propose a global quaternary structure model and a catalytic mechanism of the holoenzyme (Schoeffler & Berger, 2008). The BRD binds a DNA segment termed the 'gate-' or G-segment at the DNA-gate (Fig. 3). The N-terminal ATPase domains dimerize upon ATP binding, capturing the DNA duplex to be transported (T-segment). The T-segment is then passed through a transient break in the G-segment opened by the breakage-reunion domains, the DNA is resealed and the Tsegment released through a protein gate, the C-gate, prior to resetting of the enzyme to the

Quinolones, which target the two bacterial type II topoisomerases, exert their powerful antibacterial activity by interfering with the enzymatic reaction cycle. Specifically, they bind to the enzyme-DNA binary complex, thereby stabilizing the covalent enzyme tyrosyl-DNA phosphate ester (see also section 6 and Fig. 7). The resulting ternary complexes block DNA

DNA double helix.

Fig. 1. Chemical structures of a fluoroquinolone (left) and moxifloxacin (right). The fluorine atom, signature of fluoroquinolones, is at position 6 in the quinoline ring. In the moxifloxacin structure, oxygen atoms are coloured in red, nitrogen atoms in blue, hydrogen atoms in green and the fluor atom in purple. R1 is a cyclopropyl group, R5 is a hydrogen atom, R7 is an azabicyclo group and X8 a methyl ether group.

#### **2. Bacterial type II topoisomerases are the target of fluoroquinolones**

Topoisomerases are ubiquitous nucleic acid-dependant nanomachines essential to cell life and solve the topological problems of DNA that occur as a result of DNA manipulations during replication, transcription, recombination, chromosome segregation and condensation, maintenance of nuclear architecture and apoptosis (Champoux, 2001). They have been divided into two classes, type I and type II, according to their basic mechanism of action (Champoux, 2001; Forterre et al., 2007). Type I DNA topoisomerases introduce transient single-stranded breaks to force the passage of one DNA strand through the other, whereas type II DNA topoisomerases introduce transient double-stranded breaks to force the passage of a second DNA duplex through this broken duplex. All organisms contain at least one type I and one type II topoisomerases. Type I and type II topoisomerases can sometimes perform similar function *in vivo*, for example efficient separation of interlocked chromosomes at the end of DNA replication via their decatenation activity (Nurse et al., 2003). All type II topoisomerases can also relax both positive and negative superturns that accumulate during transcription and replication. Topoisomerases thus play an essential role for the preservation of genome stability in all life forms (Forterre et al., 2007).

Type II topoisomerases have been further sub-classified into two families, type IIA and IIB, based on evolutionary considerations (Champoux, 2001; Forterre et al., 2007). The type IIA includes bacterial DNA gyrase and topoisomerase IV, eukaryal and viral topoisomerases, whereas the type IIB includes archaeal topoisomerase VI, and their homologues in plants, a few protists, and a few bacteria (Forterre et al., 2007; Forterre & Gadelle, 2009). Type IIA and IIB topoisomerases are both formed by the association of two subunits, sharing homologous sequences and domain organization (Fig. 2). Bacterial type II topoisomerases, DNA gyrase and topoisomerase IV, consist of two subunits, GyrA or ParC and GyrB or ParE, which form the catalytic active heterotetrameric (A2B2/C2E2) complex. Subunit A consists of two domains, the N-terminal breakage-reunion domain and a carboxy-terminal domain, termed BRD and CTD, respectively (represented in blue and green respectively in Fig. 2 and 3). Subunit B consists of the ATPase domain followed by the Toprim domain (represented in yellow and red, respectively, in Fig. 2 and Fig. 3). The GyrB Toprim domain and GyrA BRD come from separate subunits and cooperatively form the enzyme catalytic core (see section

Fig. 1. Chemical structures of a fluoroquinolone (left) and moxifloxacin (right). The fluorine

moxifloxacin structure, oxygen atoms are coloured in red, nitrogen atoms in blue, hydrogen atoms in green and the fluor atom in purple. R1 is a cyclopropyl group, R5 is a hydrogen

Topoisomerases are ubiquitous nucleic acid-dependant nanomachines essential to cell life and solve the topological problems of DNA that occur as a result of DNA manipulations during replication, transcription, recombination, chromosome segregation and condensation, maintenance of nuclear architecture and apoptosis (Champoux, 2001). They have been divided into two classes, type I and type II, according to their basic mechanism of action (Champoux, 2001; Forterre et al., 2007). Type I DNA topoisomerases introduce transient single-stranded breaks to force the passage of one DNA strand through the other, whereas type II DNA topoisomerases introduce transient double-stranded breaks to force the passage of a second DNA duplex through this broken duplex. All organisms contain at least one type I and one type II topoisomerases. Type I and type II topoisomerases can sometimes perform similar function *in vivo*, for example efficient separation of interlocked chromosomes at the end of DNA replication via their decatenation activity (Nurse et al., 2003). All type II topoisomerases can also relax both positive and negative superturns that accumulate during transcription and replication. Topoisomerases thus play an essential role

atom, signature of fluoroquinolones, is at position 6 in the quinoline ring. In the

**2. Bacterial type II topoisomerases are the target of fluoroquinolones**

for the preservation of genome stability in all life forms (Forterre et al., 2007).

Type II topoisomerases have been further sub-classified into two families, type IIA and IIB, based on evolutionary considerations (Champoux, 2001; Forterre et al., 2007). The type IIA includes bacterial DNA gyrase and topoisomerase IV, eukaryal and viral topoisomerases, whereas the type IIB includes archaeal topoisomerase VI, and their homologues in plants, a few protists, and a few bacteria (Forterre et al., 2007; Forterre & Gadelle, 2009). Type IIA and IIB topoisomerases are both formed by the association of two subunits, sharing homologous sequences and domain organization (Fig. 2). Bacterial type II topoisomerases, DNA gyrase and topoisomerase IV, consist of two subunits, GyrA or ParC and GyrB or ParE, which form the catalytic active heterotetrameric (A2B2/C2E2) complex. Subunit A consists of two domains, the N-terminal breakage-reunion domain and a carboxy-terminal domain, termed BRD and CTD, respectively (represented in blue and green respectively in Fig. 2 and 3). Subunit B consists of the ATPase domain followed by the Toprim domain (represented in yellow and red, respectively, in Fig. 2 and Fig. 3). The GyrB Toprim domain and GyrA BRD come from separate subunits and cooperatively form the enzyme catalytic core (see section

atom, R7 is an azabicyclo group and X8 a methyl ether group.

5). The BRD contains the catalytic tyrosine responsible for the cleavage and religation of the DNA double helix.

Fig. 2. Schematic representation of the sequence and domain organization of type II topoisomerases found in eukaryotes, bacteria, virus and archaea. The type IIA topoisomerase family includes bacterial DNA gyrase and topoisomerase IV (Topo IV), eucaryal and viral topoisomerases (Topo II), whereas the type IIB topoisomerase family includes archaeal topoisomerase VI (Topo VI), and their homologues in plants and a few protists and bacteria. Topo IIA and Topo IIB are both formed by the association of two subunits A and B. Bacterial type IIA and type IIB topoisomerases are A2B2 heterotetramers, whereas the eukaryotic Topo II is a heterodimer, with the A and B subunits fused in a single polypeptide (Nter-B-A-Cter). The names of the four conserved domains are indicated. The winged helix domain (WHD) in the type IIB topoisomerase corresponds to the breakagereunion domain (BRD) of type IIA topoisomerase.

Crystallographic studies of individual domains show that the structure of the BRD is a heart-shaped arrangement with two dimer interfaces (Morais-Cabral et al., 1997), that the CTD displays a spiral six-bladed β-pinwheel structure (Corbett et al., 2004) and that the ATPase domain belongs to the GHKL ATPases, a broad family of enzymes with a common fold unrelated to other canonical ATP-binding folds (Brino et al., 2000) (Fig. 3). The Toprim domain interacts with the A subunit, possesses a magnesium-binding site and is essential for DNA binding (Fig. 3). Although no structure for a fully intact active type IIA topoisomerase has been determined yet, combination of structural and biochemical studies of the individual domains has lead several authors to propose a global quaternary structure model and a catalytic mechanism of the holoenzyme (Schoeffler & Berger, 2008). The BRD binds a DNA segment termed the 'gate-' or G-segment at the DNA-gate (Fig. 3). The N-terminal ATPase domains dimerize upon ATP binding, capturing the DNA duplex to be transported (T-segment). The T-segment is then passed through a transient break in the G-segment opened by the breakage-reunion domains, the DNA is resealed and the Tsegment released through a protein gate, the C-gate, prior to resetting of the enzyme to the open clamp form (Fig. 3).

Quinolones, which target the two bacterial type II topoisomerases, exert their powerful antibacterial activity by interfering with the enzymatic reaction cycle. Specifically, they bind to the enzyme-DNA binary complex, thereby stabilizing the covalent enzyme tyrosyl-DNA phosphate ester (see also section 6 and Fig. 7). The resulting ternary complexes block DNA

Quinolone Resistance in Tuberculosis Treatment: A Structural Overview 295

few bacteria such as *Treponema pallidum*, *Helicobacter pylori*, and notably *Mycobacterium tuberculosis*, the intracellular pathogen that causes tuberculosis (Cole et al., 1998) (Fig. 4). Consequently, these bacteria are unusual in producing a "single" type II topoisomerase. DNA gyrase is therefore the unique target of fluoroquinolones in these organisms, as it has

Fig. 4. Bacterial phylogenetic tree showing the distribution of species known to possess only

The direct consequence of the presence of a unique type II topoisomerase in *M. tuberculosis* has been investigated and it has been shown that *M. tuberculosis* DNA gyrase presents two specificities. First, this enzyme possesses an "hybrid activity". In addition to its normal supercoiling activity (e.g. comparable to that of *E. coli* DNA gyrase), it possesses a decatenation activity superior to that of DNA gyrase from species harboring two type II topoisomerases (Aubry et al., 2006). In addition, it shows an enhanced DNA relaxation and cleavage activities. However, the unique type II topoisomerase of *M. tuberculosis* decatenates kDNA less efficiently than a genuine topoisomerase IV, e.g., that of *S. pneumoniae*. The second specificity concerns the susceptibility to fluoroquinolones. The *M. tuberculosis* DNA gyrase is indeed naturally less susceptible to fluoroquinolones than other bacterial DNA gyrases. We have shown that three residues of the *M. tuberculosis* sequence play an important role in this natural resistance mechanism (Matrat et al., 2006). Replacing M74 in GyrA, A83 in GyrA, and R447 in GyrB of *M. tuberculosis* gyrase by their *E. coli* homologs (Ile, Ser and Lys, respectively, see residues pink underlined in Fig. 5) resulted in active enzymes as quinolone susceptible as the *E. coli* DNA gyrase. However, the question whether the

been demonstrated for *M. tuberculosis* (Mdluli & Ma, 2007).

one type II topoisomerase gene in their genome.

replication and lead to cell death (Hooper, 2003). In addition, hydrolysis of this linkage leads to the accumulation of double-stranded DNA fragments, which accounts for the bactericidal activity.

Fig. 3. Crystal structures of the individual domains of type II topoisomerases, as defined in Fig. 1. Structures of the GHKL domain (A), Toprim (B), BRD (C) and DNA gyrase CTD (D) domains. Two ATP molecules are represented in red in (A). (E) Global architecture of type IIA topoisomerases proposed based on the structures of the isolated domains adapted from Corbett & Berger (2004). Color code is the same than in Fig. 1. The dimerization interfaces of the heterotretramer constitute the three gates (N-, DNA- and C-gate), through which the DNA double helix (T-segment, represented in cyan) will be transported. The G-segment that binds the DNA-gate is represented in orange.

#### **3.** *M. tuberculosis* **DNA gyrase as the sole target of fluoroquinolones**

Bacterial genomes usually encode two type IIA enzymes, DNA gyrase and topoisomerase IV, that together help manage chromosome integrity and topology (Champoux, 2001). DNA gyrase is unique in introducing negative supercoils into DNA, an activity mediated by the CTD of its DNA binding subunit (GyrA) and is therefore responsible for the DNA unwinding at replication forks. Although closely related to DNA gyrase, topoisomerase IV has a specialized function in mediating the decatenation of interlocked daughter chromosomes (Levine et al., 1998) and relaxes positive supercoils. Particularly intriguing is that some bacteria possess in their genome only one type II topoisomerase, DNA gyrase.

Given the important role of DNA supercoiling in DNA replication, transcription, and chromosome dynamics, it is not surprising that DNA gyrase genes have been found in all bacterial genomes sequenced to date. In contrast, the topoisomerase IV gene is absent in a

replication and lead to cell death (Hooper, 2003). In addition, hydrolysis of this linkage leads to the accumulation of double-stranded DNA fragments, which accounts for the bactericidal

Fig. 3. Crystal structures of the individual domains of type II topoisomerases, as defined in Fig. 1. Structures of the GHKL domain (A), Toprim (B), BRD (C) and DNA gyrase CTD (D) domains. Two ATP molecules are represented in red in (A). (E) Global architecture of type IIA topoisomerases proposed based on the structures of the isolated domains adapted from Corbett & Berger (2004). Color code is the same than in Fig. 1. The dimerization interfaces of the heterotretramer constitute the three gates (N-, DNA- and C-gate), through which the DNA double helix (T-segment, represented in cyan) will be transported. The G-segment that

**3.** *M. tuberculosis* **DNA gyrase as the sole target of fluoroquinolones**

Bacterial genomes usually encode two type IIA enzymes, DNA gyrase and topoisomerase IV, that together help manage chromosome integrity and topology (Champoux, 2001). DNA gyrase is unique in introducing negative supercoils into DNA, an activity mediated by the CTD of its DNA binding subunit (GyrA) and is therefore responsible for the DNA unwinding at replication forks. Although closely related to DNA gyrase, topoisomerase IV has a specialized function in mediating the decatenation of interlocked daughter chromosomes (Levine et al., 1998) and relaxes positive supercoils. Particularly intriguing is that some bacteria possess in their genome only one type II topoisomerase, DNA gyrase.

Given the important role of DNA supercoiling in DNA replication, transcription, and chromosome dynamics, it is not surprising that DNA gyrase genes have been found in all bacterial genomes sequenced to date. In contrast, the topoisomerase IV gene is absent in a

binds the DNA-gate is represented in orange.

activity.

few bacteria such as *Treponema pallidum*, *Helicobacter pylori*, and notably *Mycobacterium tuberculosis*, the intracellular pathogen that causes tuberculosis (Cole et al., 1998) (Fig. 4). Consequently, these bacteria are unusual in producing a "single" type II topoisomerase. DNA gyrase is therefore the unique target of fluoroquinolones in these organisms, as it has been demonstrated for *M. tuberculosis* (Mdluli & Ma, 2007).

Fig. 4. Bacterial phylogenetic tree showing the distribution of species known to possess only one type II topoisomerase gene in their genome.

The direct consequence of the presence of a unique type II topoisomerase in *M. tuberculosis* has been investigated and it has been shown that *M. tuberculosis* DNA gyrase presents two specificities. First, this enzyme possesses an "hybrid activity". In addition to its normal supercoiling activity (e.g. comparable to that of *E. coli* DNA gyrase), it possesses a decatenation activity superior to that of DNA gyrase from species harboring two type II topoisomerases (Aubry et al., 2006). In addition, it shows an enhanced DNA relaxation and cleavage activities. However, the unique type II topoisomerase of *M. tuberculosis* decatenates kDNA less efficiently than a genuine topoisomerase IV, e.g., that of *S. pneumoniae*. The second specificity concerns the susceptibility to fluoroquinolones. The *M. tuberculosis* DNA gyrase is indeed naturally less susceptible to fluoroquinolones than other bacterial DNA gyrases. We have shown that three residues of the *M. tuberculosis* sequence play an important role in this natural resistance mechanism (Matrat et al., 2006). Replacing M74 in GyrA, A83 in GyrA, and R447 in GyrB of *M. tuberculosis* gyrase by their *E. coli* homologs (Ile, Ser and Lys, respectively, see residues pink underlined in Fig. 5) resulted in active enzymes as quinolone susceptible as the *E. coli* DNA gyrase. However, the question whether the

Quinolone Resistance in Tuberculosis Treatment: A Structural Overview 297

"single" DNA gyrases of *Treponema pallidum* or *Helicobacter pylori* display similar specificities, like increased decatenation activity or lower susceptibility to quinolones, is still

**4. The quinolone resistance determining regions and the quinolone binding**

Bacterial resistance to antibiotics typically involves drug inactivation or modification, target alteration, or decrease in drug accumulation associated with decrease in permeability and increase in efflux (Li & Nikaido, 2004). Fluoroquinolone resistance mechanisms can involve two of these three mechanisms, target alterations and overexpression of intrinsic multidrug resistance (MDR) efflux pumps (Hooper, 2002). However, resistance to fluoroquinolone is mainly due to target modification, e.g. mutations in the DNA gyrase and/or topoisomerase IV genes. Mutations conferring bacterial resistance to quinolones occur in two short discrete segments termed the quinolone resistance-determining regions (QRDR) in the breakagereunion domain of subunit A (QRDR-A) and more rarely in the Toprim domain of subunit B

Fig. 6. Schematic representation of the *M. tuberculosis* DNA gyrase sequence. Subunit A contains the breakage-reunion domain (BRD, blue) and the C-terminal domain (CTD, green). The subunit B contains the ATPase domain (yellow) and the Toprim domain (red). Localization of the QRDR is indicated in pink and light blue. The color code is the same than in Fig. 2 and 3. The approximate lengths of the QRDRs are indicated (30 and 40 aa for the QRDR-A and -B, respectively). The residue ranges of the QRDR-A and -B are indicated for

Whereas residues of the QRDR-A have been known for a while that they are spatially close to the catalytic tyrosine responsible for the double-stranded DNA cleavage, residues of the QRDR-B were only hypothetically thought to be involved in the quinolone-binding pocket (Heddle & Maxwell, 2002). This ambiguity was brought about by the two first structures of the catalytic core (an homodimer formed by two single polypeptides composed of the Toprim domain and the BRD) of the yeast topoisomerase II (Fass et al., 1999; Berger & Gamblin, 1996). In these structures, which correspond to two different conformational arrangements of the catalytic core, the two QRDRs are too far from each other to form a

the *M. tuberculosis* and the *E. coli* DNA gyrase sequences.

unique binding pocket for the quinolone molecule (Fig. 7).

open.

**pocket**

(QRDR-B) (Hooper, 1999) (Fig. 6).

Fig. 5. Two parts of the multiple alignment of subunit A and B, respectively, centred on the quinolone resistance determining regions (QRDR, see section 4 for definition), after Matrat et al. (2008). Residues highlighted in blue underline the most important residues shown to be involved in acquired resistance to fluoroquinolones. Residues highlighted in pink underline residues of the *M. tuberculosis* DNA gyrase sequence involved in the natural resistance mechanism. The numbering at the top of the alignments corresponds to the *E. coli* DNA gyrase numbering. The values of MIC50 and IC50 are given for ofloxacin when known.

Fig. 5. Two parts of the multiple alignment of subunit A and B, respectively, centred on the quinolone resistance determining regions (QRDR, see section 4 for definition), after Matrat et al. (2008). Residues highlighted in blue underline the most important residues shown to be involved in acquired resistance to fluoroquinolones. Residues highlighted in pink underline residues of the *M. tuberculosis* DNA gyrase sequence involved in the natural resistance mechanism. The numbering at the top of the alignments corresponds to the *E. coli* DNA gyrase numbering. The values of MIC50 and IC50 are given for ofloxacin when known. "single" DNA gyrases of *Treponema pallidum* or *Helicobacter pylori* display similar specificities, like increased decatenation activity or lower susceptibility to quinolones, is still open.

## **4. The quinolone resistance determining regions and the quinolone binding pocket**

Bacterial resistance to antibiotics typically involves drug inactivation or modification, target alteration, or decrease in drug accumulation associated with decrease in permeability and increase in efflux (Li & Nikaido, 2004). Fluoroquinolone resistance mechanisms can involve two of these three mechanisms, target alterations and overexpression of intrinsic multidrug resistance (MDR) efflux pumps (Hooper, 2002). However, resistance to fluoroquinolone is mainly due to target modification, e.g. mutations in the DNA gyrase and/or topoisomerase IV genes. Mutations conferring bacterial resistance to quinolones occur in two short discrete segments termed the quinolone resistance-determining regions (QRDR) in the breakagereunion domain of subunit A (QRDR-A) and more rarely in the Toprim domain of subunit B (QRDR-B) (Hooper, 1999) (Fig. 6).

Fig. 6. Schematic representation of the *M. tuberculosis* DNA gyrase sequence. Subunit A contains the breakage-reunion domain (BRD, blue) and the C-terminal domain (CTD, green). The subunit B contains the ATPase domain (yellow) and the Toprim domain (red). Localization of the QRDR is indicated in pink and light blue. The color code is the same than in Fig. 2 and 3. The approximate lengths of the QRDRs are indicated (30 and 40 aa for the QRDR-A and -B, respectively). The residue ranges of the QRDR-A and -B are indicated for the *M. tuberculosis* and the *E. coli* DNA gyrase sequences.

Whereas residues of the QRDR-A have been known for a while that they are spatially close to the catalytic tyrosine responsible for the double-stranded DNA cleavage, residues of the QRDR-B were only hypothetically thought to be involved in the quinolone-binding pocket (Heddle & Maxwell, 2002). This ambiguity was brought about by the two first structures of the catalytic core (an homodimer formed by two single polypeptides composed of the Toprim domain and the BRD) of the yeast topoisomerase II (Fass et al., 1999; Berger & Gamblin, 1996). In these structures, which correspond to two different conformational arrangements of the catalytic core, the two QRDRs are too far from each other to form a unique binding pocket for the quinolone molecule (Fig. 7).

Quinolone Resistance in Tuberculosis Treatment: A Structural Overview 299

the oxygen of the tyrosine and the phosphor atom of a phosphate group (Fig. 8). In addition, this structure also confirmed that the QRDR-A corresponds to the DNA-binding domain of the subunit A often referred to as a CAP-like domain constituted by a helix-turn-helix motif (H3 and H4). The QRDR-B corresponds to the DNA-binding region of subunit B that involves a helix and a long loop (see section 5). However, the main conclusion of this work was that the conformational arrangement observed in this structure brings together the two QRDRs to form a unique quinolone-binding pocket (see section 6) at the interface formed by

Fig. 8. Crystal structure of the *S. pneumoniae* topoisomerase IV catalytic core that comprises the GyrB Toprim domain and GyrA BRD in complex with DNA and moxifloxacin. The

oligonucleotide in orange and the moxifloxacin in green. (Top) Side and top views of the molecular surface of the catalytic core. (Bottom) Close view of the catalytic tyrosine (represented in blue sticks) before and after formation of the covalent link between the enzyme and DNA (the phosphodiester bond is indicated by a black arrow). Residues of the

Toprim domain is represented in red, the BRD in blue, the 35 base pairs DNA

Toprim domain chelating the magnesium ion are represented in red sticks.

the DNA, the BRD and the Toprim domain (Fig. 8).

Fig. 7. Side view (top left and right) and top view (bottom left and right) of the catalytic core of the yeast topoisomerase II, composed of the Toprim domain (represented in red) and the breakage-reunion domain (BRD, represented in blue). (Left) "Open" conformation of the catalytic core (PDB code 1BGW, Berger et al., 1996). The DNA-gate is open and the C-gate is closed (for definition of DNA- and C-gate, see Fig. 3). The QRDR-B (represented in cyan) are close together but distant from the QRDR-A (represented in purple). The QRDR-A includes the two DNA-binding helices, H3 and H4. (Right) "Intermediate" conformation of the catalytic core (PDB code 1BJT, Fass et al., 1999). The DNA-gate is slightly open and the Cgate is closed. The QRDR-B are more distant from each other and are closer to the QRDR-A, but still do not form a unique quinolone-binding pocket.

Involvement of QRDR-B residues in the quinolone-binding pocket was definitely confirmed in 2009 with the first structure of the *S. pneumoniae* topoisomerase IV Toprim and breakagereunion domains in complex with DNA and moxifloxacin (Lapogonov et al., 2009). This structure highlighted the fact that the GyrA BRD and the GyrB-Toprim domain form the catalytic core complex, where the double-stranded DNA interacts at the interface of the two domains (Fig. 8). This structure brought for the first time the structural evidence of the covalent link formed between the DNA and the enzyme, a phosphodiester bond between

Fig. 7. Side view (top left and right) and top view (bottom left and right) of the catalytic core of the yeast topoisomerase II, composed of the Toprim domain (represented in red) and the breakage-reunion domain (BRD, represented in blue). (Left) "Open" conformation of the catalytic core (PDB code 1BGW, Berger et al., 1996). The DNA-gate is open and the C-gate is closed (for definition of DNA- and C-gate, see Fig. 3). The QRDR-B (represented in cyan) are close together but distant from the QRDR-A (represented in purple). The QRDR-A includes the two DNA-binding helices, H3 and H4. (Right) "Intermediate" conformation of the catalytic core (PDB code 1BJT, Fass et al., 1999). The DNA-gate is slightly open and the Cgate is closed. The QRDR-B are more distant from each other and are closer to the QRDR-A,

Involvement of QRDR-B residues in the quinolone-binding pocket was definitely confirmed in 2009 with the first structure of the *S. pneumoniae* topoisomerase IV Toprim and breakagereunion domains in complex with DNA and moxifloxacin (Lapogonov et al., 2009). This structure highlighted the fact that the GyrA BRD and the GyrB-Toprim domain form the catalytic core complex, where the double-stranded DNA interacts at the interface of the two domains (Fig. 8). This structure brought for the first time the structural evidence of the covalent link formed between the DNA and the enzyme, a phosphodiester bond between

but still do not form a unique quinolone-binding pocket.

the oxygen of the tyrosine and the phosphor atom of a phosphate group (Fig. 8). In addition, this structure also confirmed that the QRDR-A corresponds to the DNA-binding domain of the subunit A often referred to as a CAP-like domain constituted by a helix-turn-helix motif (H3 and H4). The QRDR-B corresponds to the DNA-binding region of subunit B that involves a helix and a long loop (see section 5). However, the main conclusion of this work was that the conformational arrangement observed in this structure brings together the two QRDRs to form a unique quinolone-binding pocket (see section 6) at the interface formed by the DNA, the BRD and the Toprim domain (Fig. 8).

Fig. 8. Crystal structure of the *S. pneumoniae* topoisomerase IV catalytic core that comprises the GyrB Toprim domain and GyrA BRD in complex with DNA and moxifloxacin. The Toprim domain is represented in red, the BRD in blue, the 35 base pairs DNA oligonucleotide in orange and the moxifloxacin in green. (Top) Side and top views of the molecular surface of the catalytic core. (Bottom) Close view of the catalytic tyrosine (represented in blue sticks) before and after formation of the covalent link between the enzyme and DNA (the phosphodiester bond is indicated by a black arrow). Residues of the Toprim domain chelating the magnesium ion are represented in red sticks.

Quinolone Resistance in Tuberculosis Treatment: A Structural Overview 301

moxifloxacin. In both the crystal structure of the *S. pneumoniae* topoisomerase IV and the modeled *M. tuberculosis* DNA gyrase catalytic cores, the fluoroquinolone molecules are bound at the ternary interface formed by the DNA, the BRD and the Toprim domain (Fig. 8 and 10). The general mode of action known for quinolones, common to DNA gyrase and topoisomerase IV, was the inhibition of the broken DNA religation. The way how these molecules are responsible for this inhibition has been clarified by the structure. Whereas quinolones are not DNA intercalators, the fluoroquinolone molecule is intercalated in the ternary complex between the dinucleotide step where the covalent bond between the DNA and the enzyme (with the catalytic tyrosine) is formed (Fig. 8 and 10). As the catalytic core is dimeric, two fluoroquinolones molecules are bound spaced by four base pairs (Fig. 10). The consequence of this intercalation is that the O3'-P religation is structurally impossible

Fig. 10. Structure of the *M. tuberculosis* DNA gyrase catalytic core in complex with DNA and moxifloxacin. Side view (top left) and top view (top right) of the molecular surface of the catalytic core. The Toprim domain is represented in red, the BRD in blue, the 35 base pairs DNA oligonucleotide in orange and the moxifloxacin in green. (Bottom left) Close view of the structure of the intercalated moxifloxacin (magenta) in the broken DNA double helix (green). The catalytic tyrosine (Y129 in the *M. tuberculosis* DNA gyrase sequence) is shown in green outside the DNA helix. (Bottom right) Close view of both moxifloxacin molecules in the broken DNA showing the four base pairs in between the two bound fluoroquinolones. Both catalytic tyrosines of each monomer are shown in green in the DNA major groove.

because both atoms are too far away from each other (more than 9 Å) (Fig. 10).

## **5. Structural studies of the** *M. tuberculosis* **DNA gyrase catalytic core**

We recently solved the crystal structures of two domains of the *M. tuberculosis* DNA gyrase, the GyrB Toprim (TopBK) and the GyrA breakage-reunion (GA57BK) domains (Piton et al., 2009; 2010). These two domains form the DNA gyrase catalytic core that corresponds to the smallest entity able to catalyse DNA cleavage (Piton et al., 2010).

We solved two high resolution structures of the Toprim domain to 2.1 and 1.95 Å resolution, respectively. These two structures display two different conformations of the magnesium binding site. The QRDR-B is constituted by residues 500 to 538 (461-499 in the PDB numbering). Part of this region is disordered and corresponds to a loop that has been named DNA-binding loop (DBL) because it is folded in the presence of DNA (Fig. 9). We obtained four crystal forms of the breakage-reunion domain, diffracting from 4.2 to 2.7 Å resolution depending on the crystal form (Piton et al., 2009). This was the first structure determination report of the BRD of a DNA gyrase from a species containing one unique type II topoisomerase. The structure of this domain is unexpectedly very similar to other structures of BRD solved for bacterial type II topoisomerases, DNA gyrase and topoisomerase IV (Morais-Cabral et al., 1997; Carr et al., 2007). It displays the typical heart-like shaped structure with a hole of 30 Å diameter at the center allowing the passage of the DNA from the DNA-gate to the C-gate (Fig. 9). The QRDR-A is localised at the DNA-gate (shown in purple in Fig. 9).

Fig. 9. Crystals (top) and crystal structures (down) of the GyrB Toprim (TopBK, PDB code 3IG0, 242 residues) domain represented in red (left) and the GyrA breakage-reunion (BRD, GA57BK, PDB code 3IFZ, 508 residues per monomer) represented in blue (right). The QRDR-B is represented in cyan in the TopBK structure (residues 500-538), the QRDR-A is represented in purple in the GA57BK structure (residues 74-101).

Using the two crystal structures we determined, the GyrA BRD (GA57BK) and the GyrB Toprim (TopBK) domains, we performed the modeling of the *M. tuberculosis* catalytic core in complex with a 35 base pairs DNA oligonucleotide and the most used fluoroquinolone,

We recently solved the crystal structures of two domains of the *M. tuberculosis* DNA gyrase, the GyrB Toprim (TopBK) and the GyrA breakage-reunion (GA57BK) domains (Piton et al., 2009; 2010). These two domains form the DNA gyrase catalytic core that corresponds to the

We solved two high resolution structures of the Toprim domain to 2.1 and 1.95 Å resolution, respectively. These two structures display two different conformations of the magnesium binding site. The QRDR-B is constituted by residues 500 to 538 (461-499 in the PDB numbering). Part of this region is disordered and corresponds to a loop that has been named DNA-binding loop (DBL) because it is folded in the presence of DNA (Fig. 9). We obtained four crystal forms of the breakage-reunion domain, diffracting from 4.2 to 2.7 Å resolution depending on the crystal form (Piton et al., 2009). This was the first structure determination report of the BRD of a DNA gyrase from a species containing one unique type II topoisomerase. The structure of this domain is unexpectedly very similar to other structures of BRD solved for bacterial type II topoisomerases, DNA gyrase and topoisomerase IV (Morais-Cabral et al., 1997; Carr et al., 2007). It displays the typical heart-like shaped structure with a hole of 30 Å diameter at the center allowing the passage of the DNA from the DNA-gate to the C-gate (Fig. 9). The QRDR-A is localised at the DNA-gate (shown in

Fig. 9. Crystals (top) and crystal structures (down) of the GyrB Toprim (TopBK, PDB code 3IG0, 242 residues) domain represented in red (left) and the GyrA breakage-reunion (BRD, GA57BK, PDB code 3IFZ, 508 residues per monomer) represented in blue (right). The QRDR-B is represented in cyan in the TopBK structure (residues 500-538), the QRDR-A is

Using the two crystal structures we determined, the GyrA BRD (GA57BK) and the GyrB Toprim (TopBK) domains, we performed the modeling of the *M. tuberculosis* catalytic core in complex with a 35 base pairs DNA oligonucleotide and the most used fluoroquinolone,

represented in purple in the GA57BK structure (residues 74-101).

**5. Structural studies of the** *M. tuberculosis* **DNA gyrase catalytic core**

smallest entity able to catalyse DNA cleavage (Piton et al., 2010).

purple in Fig. 9).

moxifloxacin. In both the crystal structure of the *S. pneumoniae* topoisomerase IV and the modeled *M. tuberculosis* DNA gyrase catalytic cores, the fluoroquinolone molecules are bound at the ternary interface formed by the DNA, the BRD and the Toprim domain (Fig. 8 and 10). The general mode of action known for quinolones, common to DNA gyrase and topoisomerase IV, was the inhibition of the broken DNA religation. The way how these molecules are responsible for this inhibition has been clarified by the structure. Whereas quinolones are not DNA intercalators, the fluoroquinolone molecule is intercalated in the ternary complex between the dinucleotide step where the covalent bond between the DNA and the enzyme (with the catalytic tyrosine) is formed (Fig. 8 and 10). As the catalytic core is dimeric, two fluoroquinolones molecules are bound spaced by four base pairs (Fig. 10). The consequence of this intercalation is that the O3'-P religation is structurally impossible because both atoms are too far away from each other (more than 9 Å) (Fig. 10).

Fig. 10. Structure of the *M. tuberculosis* DNA gyrase catalytic core in complex with DNA and moxifloxacin. Side view (top left) and top view (top right) of the molecular surface of the catalytic core. The Toprim domain is represented in red, the BRD in blue, the 35 base pairs DNA oligonucleotide in orange and the moxifloxacin in green. (Bottom left) Close view of the structure of the intercalated moxifloxacin (magenta) in the broken DNA double helix (green). The catalytic tyrosine (Y129 in the *M. tuberculosis* DNA gyrase sequence) is shown in green outside the DNA helix. (Bottom right) Close view of both moxifloxacin molecules in the broken DNA showing the four base pairs in between the two bound fluoroquinolones. Both catalytic tyrosines of each monomer are shown in green in the DNA major groove.

Quinolone Resistance in Tuberculosis Treatment: A Structural Overview 303

pocket too small to bind moxifloxacin (Fig. 12). In the QRDR-B, the side chain of residue R521, an amino acid located in the Toprim DBL, points towards the DNA minor groove and forms a "flap" that blocks the fluoroquinolone in the pocket through contact with the R7 group (Fig. 11). The increase of susceptibility to fluoroquinolones by the substitution of R521 to lysine can be explained by the lower energy cost when moving an arginine than a lysine in the DNA minor groove (Rohs et al., 2010). This means that the "flap" created by this amino acid, would find it easier to open up and destabilize the fluoroquinolone, when this

Fig. 11. (Top) Schematic representation of the *M. tuberculosis* DNA gyrase sequence. Subunit A contains the breakage-reunion domain (BRD, blue) and the C-terminal domain (CTD, green). The subunit B contains the ATPase domain (yellow) and the Toprim domain (red). *End of legend*. Localization of the QRDR is indicated in pink and light blue. (Bottom left) Top view of the molecular surface representation of the M. *tuberculosis* DNA gyrase catalytic core in complex with DNA and moxifloxacin showing the QBP (color code is the same than in Fig. 10). The QRDR-A and -B are represented in pink and light blue, respectively. (Right) Close view of the quinolone-binding pocket. The DNA-protein complex is represented in transparent molecular surface and moxifloxacin in sticks (color code is the same than in Fig. 10). The residues of the QRDR-B (Toprim) belonging to the QBP are indicated in pink, purple and yellow. Resides A90 of the QRDR-B is represented in light green in the

residue is an arginine rather than a lysine.

background of the pocket.

#### **6. Fluoroquinolone resistance in** *M. tuberculosis*

Fluoroquinolone resistance in *M. tuberculosis* is defined by resistance to ofloxacin (ability of the bacilli to grow on medium containing 2 mg/l ofloxacin (WHO, 2001) and is mainly due to mutations in DNA gyrase in the QRDRs (see section 4) (Takiff et al., 1994). Mutations within the QRDRs are responsible of at least 75% of fluoroquinolone resistance in *M. tuberculosis*, affecting most commonly the GyrA subunit (G88A, A90V, S91P, D94G, H, N, Y) but also the GyrB subunit (Aubry et al., 2006b; Veziris et al., 2007; Siddiqi et al., 2002; Feuerriegel et al., 2009). Modeling of the *M. tuberculosis* DNA gyrase catalytic core allowed to clearly establish that QRDR residues of both subunits are spatially close and form the quinolone-binding pocket (QBP) (Fig. 11). In this pocket, the fluoroquinolone is maintained on one side by three residues of the QRDR-A, G88, D89 and A90, that are in close contact with the quinolone conserved carboxylic function (R3 group, Fig. 1 and 11), and on the other side by five residues of the QRDR-B, D500, R521, N538, T539 and E540 in close contact with the R1, R7 and R8 groups of the quinolone (Fig. 1 and 11).  The model shows that the geometry of the QBP is crucial for the recognition and the stability of fluoroquinolone in the pocket. Any modification of the amino acids belonging to the QBP leads to either a direct or indirect change of the geometry of the pocket (Fig. 11). The pocket becomes too large to stabilize the fluoroquinolone, when the side chain of the modified amino acid becomes smaller. In contrast, the pocket becomes too small to fix the fluoroquinolone, when the side chain of the modified amino acid is bulkier. An indirect effect on the geometry of the pocket is observed when amino acid substitutions are localised in the H4 helix that interacts with the major groove of the DNA. As DNA is also part of the QBP (Fig. 11), modification of the structure of the DNA will modify the geometry of the pocket, and can lead to destabilisation of the fluoroquinolones in the pocket. This mechanism could explain why substitutions of the amino acid D94 (position mostly found in *M. tuberculosis* strains resistant to quinolones) have paradoxical effects, e.g. the substitution by an amino acid with a smaller side chain, such as alanine or glycine, increases the resistance to the same level than the substitutions by amino acids with bulky side chains.

Resistance to fluoroquinolones of *M. tuberculosis* DNA gyrase results from two mechanisms. On one hand, the lower natural affinity of the *M. tuberculosis* DNA gyrase catalytic core for fluoroquinolones is responsible for "intrinsic resistance" and is attributable to the amino acid nature at three positions, 81 and 90 in the QRDR-A and 521 in the QRDR-B (see section 3). On the other hand, DNA gyrase modifications found in the QRDRs of DNA gyrases of clinical *M. tuberculosis* strains are responsible for "acquired resistance". Interestingly, amino acid at two positions in the QBP play a crucial role in both natural and acquired resistance, position 90 in the QRDR-A, which is an alanine in the *M. tuberculosis* sequence and position 521, an arginine in the *M. tuberculosis* QRDR-B. Both amino acid are essential for the shape of the QBP and, as a result, for the binding of quinolone. For example, the substitution of residue A90 in the wild-type *M. tuberculosis* DNA gyrase to S90 generates a pocket better adapted to the size of moxifloxacin (Fig. 12), increasing the susceptibility to this inhibitor. In contrast, the A90V substitution, found in some strains resistant to moxifloxacin, generates a

Fluoroquinolone resistance in *M. tuberculosis* is defined by resistance to ofloxacin (ability of the bacilli to grow on medium containing 2 mg/l ofloxacin (WHO, 2001) and is mainly due to mutations in DNA gyrase in the QRDRs (see section 4) (Takiff et al., 1994). Mutations within the QRDRs are responsible of at least 75% of fluoroquinolone resistance in *M. tuberculosis*, affecting most commonly the GyrA subunit (G88A, A90V, S91P, D94G, H, N, Y) but also the GyrB subunit (Aubry et al., 2006b; Veziris et al., 2007; Siddiqi et al., 2002; Feuerriegel et al., 2009). Modeling of the *M. tuberculosis* DNA gyrase catalytic core allowed to clearly establish that QRDR residues of both subunits are spatially close and form the quinolone-binding pocket (QBP) (Fig. 11). In this pocket, the fluoroquinolone is maintained on one side by three residues of the QRDR-A, G88, D89 and A90, that are in close contact with the quinolone conserved carboxylic function (R3 group, Fig. 1 and 11), and on the other side by five residues of the QRDR-B, D500, R521, N538, T539 and E540 in close contact with the R1, R7 and R8 groups of the quinolone (Fig. 1 and 11).  The model shows that the geometry of the QBP is crucial for the recognition and the stability of fluoroquinolone in the pocket. Any modification of the amino acids belonging to the QBP leads to either a direct or indirect change of the geometry of the pocket (Fig. 11). The pocket becomes too large to stabilize the fluoroquinolone, when the side chain of the modified amino acid becomes smaller. In contrast, the pocket becomes too small to fix the fluoroquinolone, when the side chain of the modified amino acid is bulkier. An indirect effect on the geometry of the pocket is observed when amino acid substitutions are localised in the H4 helix that interacts with the major groove of the DNA. As DNA is also part of the QBP (Fig. 11), modification of the structure of the DNA will modify the geometry of the pocket, and can lead to destabilisation of the fluoroquinolones in the pocket. This mechanism could explain why substitutions of the amino acid D94 (position mostly found in *M. tuberculosis* strains resistant to quinolones) have paradoxical effects, e.g. the substitution by an amino acid with a smaller side chain, such as alanine or glycine, increases the resistance to the same level than the substitutions by

Resistance to fluoroquinolones of *M. tuberculosis* DNA gyrase results from two mechanisms. On one hand, the lower natural affinity of the *M. tuberculosis* DNA gyrase catalytic core for fluoroquinolones is responsible for "intrinsic resistance" and is attributable to the amino acid nature at three positions, 81 and 90 in the QRDR-A and 521 in the QRDR-B (see section 3). On the other hand, DNA gyrase modifications found in the QRDRs of DNA gyrases of clinical *M. tuberculosis* strains are responsible for "acquired resistance". Interestingly, amino acid at two positions in the QBP play a crucial role in both natural and acquired resistance, position 90 in the QRDR-A, which is an alanine in the *M. tuberculosis* sequence and position 521, an arginine in the *M. tuberculosis* QRDR-B. Both amino acid are essential for the shape of the QBP and, as a result, for the binding of quinolone. For example, the substitution of residue A90 in the wild-type *M. tuberculosis* DNA gyrase to S90 generates a pocket better adapted to the size of moxifloxacin (Fig. 12), increasing the susceptibility to this inhibitor. In contrast, the A90V substitution, found in some strains resistant to moxifloxacin, generates a

**6. Fluoroquinolone resistance in** *M. tuberculosis*

amino acids with bulky side chains.

pocket too small to bind moxifloxacin (Fig. 12). In the QRDR-B, the side chain of residue R521, an amino acid located in the Toprim DBL, points towards the DNA minor groove and forms a "flap" that blocks the fluoroquinolone in the pocket through contact with the R7 group (Fig. 11). The increase of susceptibility to fluoroquinolones by the substitution of R521 to lysine can be explained by the lower energy cost when moving an arginine than a lysine in the DNA minor groove (Rohs et al., 2010). This means that the "flap" created by this amino acid, would find it easier to open up and destabilize the fluoroquinolone, when this residue is an arginine rather than a lysine.

Fig. 11. (Top) Schematic representation of the *M. tuberculosis* DNA gyrase sequence. Subunit A contains the breakage-reunion domain (BRD, blue) and the C-terminal domain (CTD, green). The subunit B contains the ATPase domain (yellow) and the Toprim domain (red). *End of legend*. Localization of the QRDR is indicated in pink and light blue. (Bottom left) Top view of the molecular surface representation of the M. *tuberculosis* DNA gyrase catalytic core in complex with DNA and moxifloxacin showing the QBP (color code is the same than in Fig. 10). The QRDR-A and -B are represented in pink and light blue, respectively. (Right) Close view of the quinolone-binding pocket. The DNA-protein complex is represented in transparent molecular surface and moxifloxacin in sticks (color code is the same than in Fig. 10). The residues of the QRDR-B (Toprim) belonging to the QBP are indicated in pink, purple and yellow. Resides A90 of the QRDR-B is represented in light green in the background of the pocket.

Quinolone Resistance in Tuberculosis Treatment: A Structural Overview 305

We greatly acknowledge Dr Jérémie Piton for having provided figures 8, 11 and 12. We are especially grateful to Dr Jérémie Piton and Dr Stéphanie Petrella for helpful discussions. We

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**9. References**

Fig. 12. Effect of the substitution of A90 (QRDR-A) on the geometry of the quinolonebinding pocket. (Left) Quinolone-binding pocket of the wild-type *M. tuberculosis* DNA gyrase. The A90 is coloured in yellow. (Middle) Substitution of A90 to serine (S90 is represented in green). (Right) Substitution of A90 to valine (V90 is represented in magenta).

## **7. Conclusion**

This chapter has described the structural insights into the bacterial type II topoisomerases, the targets of quinolones and into the quinolone resistance mechanism, discussed in the context of the enzyme modifications, e.g. amino acid substitutions, described in the literature that are implicated in quinolone resistance. It has shown that the tridimensional structure of the *M. tuberculosis* catalytic core in complex with DNA and a quinolone molecule is a powerful tool to better understand the relationships between the sequence, the structure and the resistance phenotype in *M. tuberculosis* DNA gyrase.

Knowledge of structural data has applications both in the diagnosis of resistance in *M. tuberculosis* but also in tuberculosis treatment. This powerfulness consists in four essential aspects. It strongly contributes to (1) the analyses of the results obtained by molecular studies since the variety of new mutations, especially in the GyrB subunit, is increasing, (2) the discrimination between substitutions identified in fluoroquinoloneresistant clinical strains that are implicated or not in fluoroquinolone resistance (Pantel et al., 2011), (3) the prediction of the effect on susceptibility to quinolones of new undescribed mutations, and finally (4) to the design of new quinolones, as quinolones used nowadays for tuberculosis treatment have been designed before structural data on the *M. tuberculosis* DNA gyrase catalytic core were available.

## **8. Acknowledgment**

We greatly acknowledge Dr Jérémie Piton for having provided figures 8, 11 and 12. We are especially grateful to Dr Jérémie Piton and Dr Stéphanie Petrella for helpful discussions. We thank Dr Olivier Poch for careful reading of the manuscript.

#### **9. References**

304 Understanding Tuberculosis – New Approaches to Fighting Against Drug Resistance

Fig. 12. Effect of the substitution of A90 (QRDR-A) on the geometry of the quinolonebinding pocket. (Left) Quinolone-binding pocket of the wild-type *M. tuberculosis* DNA gyrase. The A90 is coloured in yellow. (Middle) Substitution of A90 to serine (S90 is

structure and the resistance phenotype in *M. tuberculosis* DNA gyrase.

DNA gyrase catalytic core were available.

**7. Conclusion**

represented in green). (Right) Substitution of A90 to valine (V90 is represented in magenta).

This chapter has described the structural insights into the bacterial type II topoisomerases, the targets of quinolones and into the quinolone resistance mechanism, discussed in the context of the enzyme modifications, e.g. amino acid substitutions, described in the literature that are implicated in quinolone resistance. It has shown that the tridimensional structure of the *M. tuberculosis* catalytic core in complex with DNA and a quinolone molecule is a powerful tool to better understand the relationships between the sequence, the

Knowledge of structural data has applications both in the diagnosis of resistance in *M. tuberculosis* but also in tuberculosis treatment. This powerfulness consists in four essential aspects. It strongly contributes to (1) the analyses of the results obtained by molecular studies since the variety of new mutations, especially in the GyrB subunit, is increasing, (2) the discrimination between substitutions identified in fluoroquinoloneresistant clinical strains that are implicated or not in fluoroquinolone resistance (Pantel et al., 2011), (3) the prediction of the effect on susceptibility to quinolones of new undescribed mutations, and finally (4) to the design of new quinolones, as quinolones used nowadays for tuberculosis treatment have been designed before structural data on the *M. tuberculosis*

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**14**

**Antimycobacterial Activity Some**

**Different Lamiaceae Plant Extracts**

*Mycobacterium tuberculosis* is a pathogenic bacteria species of the genus *Mycobacterium*, first discovered in 1882 by Robert Koch, which causes tuberculosis (TB) (Ryan & Ray, 2004). The disease is characterized by symptoms such as sepsis, septic shock, multiple organ failure (Muckart & Bhagwanjee, 1997). It may spread to the central nervous system and cause TB meningitis, intracranial tuberculomas, or abscesses (Harisinghani et al., 2000; Hwang et al.,

After the late 1980s, tuberculosis morbidity and mortality rates became a major health problem for industrialized countries (Raviglione et al., 1995; Heym & Cole, 1997). Multidrug-resistant tuberculosis (MDR TB) and extensively drug resistant tuberculosis (XDR TB) has become a common phenomenon, which cause drugs to be ineffective. MDR-TB results from either primary infection or may develop in the course of a patient's treatment. MDR TB is resistant to at least two first-line anti-TB drugs, isoniazid (INH) and rifampicin (RIF), which are most powerful anti-TB drugs; XDR TB is resistant to INH and RIF, plus fluoroquinolone and at least one of three injectable second-line drugs such as capreomycin, kanamycin, and amikacin. Treatment of XDR-TB is not possible by first-line anti-TB drugs, which are less effective, expensive and toxic; in addition treatment takes two

Mycobacteria are resistant to most common antibiotics and chemotherapeutic agents due to the mycobacterial cell wall composition of bacterial peptidoglycans (Slayden & Barry, 2000; Lee et al., 1996; Brennan et al., 1995), a lipophilic layer of long-chain fatty acids, and mycolic acids (Barry et al., 1998). The rich lipids of the cell wall has an important role in their virulence (Murray, Rosenthal and Pfaller, 2005). This structure provides a highly hydrophobic and efficient barrier to antibiotics and chemotherapeutic agents (Jarlier & Nikaido 1994). Thus, this cell wall composition restricts the choice of drug treatment. Compounds capable of blocking efflux pumps so that antibiotics can gain access to their targets are of obvious importance (Viveiros et al, 2003). Increased activity of existing efflux

**1. Introduction**

years or more (WHO, 2011a; WHO, 2011b).

2010).

**Containing Flavonoids and Other**

Tulin Askun, Gulendam Tumen, Fatih Satil,

**Phenolic Compounds**

*Balikesir University*

*Turkey*

Seyma Modanlioglu and Onur Yalcin


**Comparative Effect of Mo and Cr on Microstructure and Mechanical Properties in NbV-Microalloyed Bainitic Steels**

#### **Andrii Kostryzhev 1,\*, Navjeet Singh 1, Liang Chen 2, Chris Killmore 3 and Elena Pereloma 2,4**

Received: 4 January 2018; Accepted: 13 February 2018; Published: 16 February 2018

**Abstract:** Steel product markets require the rolled stock with further increasing mechanical properties and simultaneously decreasing price. The steel cost can be reduced via decreasing the microalloying elements contents, although this decrease may undermine the mechanical properties. Multi-element microalloying with minor additions is the route to optimise steel composition and keep the properties high. However, this requires deep understanding of mutual effects of elements on each other's performance with respect to the development of microstructure and mechanical properties. This knowledge is insufficient at the moment. In the present work we investigate the microstructure and mechanical properties of bainitic steels microalloyed with Cr, Mo, Nb and V. Comparison of 0.2 wt. % Mo and Cr additions has shown a more pronounced effect of Mo on precipitation than on phase balance. Superior strength of the MoNbV-steel originated from the strong solid solution strengthening effect. Superior ductility of the CrNbV-steel corresponded to the more pronounced precipitation in this steel. Nature of these mechanisms is discussed.

**Keywords:** steel; thermomechanical processing; microstructure characterisation; mechanical properties; molybdenum

#### **1. Introduction**

In high strength low alloyed steels Mo is well known to provide phase balance strengthening, via facilitating the bainite transformation [1–5], and solid solution strengthening [6–9]. It can decrease the rate of dynamic recrystallization of austenite [10–12], which may lead to grain refinement. Sometimes Mo can contribute to precipitation strengthening through formation of Mo-rich carbides [13–16]. Although its main effect on precipitation is via the increase in solubility of Ti [17,18] and Nb [10] in austenite, resulting in decreased sizes and increased number densities of Ti- and Nb-rich particles [19–23], which are essential for the precipitation strengthening from Ti- and Nb-rich particles.

Similar to Mo, Cr facilitates the bainite transformation [24–26], may precipitate in complex Cr-rich carbides [27–29], and increases solubility of Ti [30,31] and Nb [30,32,33]. In particular, Cr was observed delaying Fe3C precipitation in low carbon steel [34]. However, the solid solution strengthening effect of Cr is ~6 times weaker than this of Mo [35], and, therefore, Cr is less affective in retarding recrystallization [36,37].

Amongst the published data, effects of Mo and Cr in multi-microalloyed steels, in particular containing V, are rarely reported. In 0.042C-0.3Mo-1.0Cr-0.08V steel coiled in the temperature range of 180–530 ◦C, Hutchinson et al. [38] observed bainitic microstructures with average ferrite grain size of 3 μm. Increased to 640–770 MPa proof stress was suggested to originate mainly from high dislocation density in bainite and, in particular, dislocation pinning by V(C,N) precipitates. In another work, Kong et al. [39] investigated mechanical properties of 0.064C-0.22Mo-0.21Cr-0.031Nb-0.031V steel thermomechanically processed in a temperature range of 1150–800 ◦C and cooled at the rate of 20–30 ◦C/s to 430–550 ◦C finish cooling temperature. The yield stress in the range of 530–710 MPa was attributed to the narrow width of bainitic ferrite lath (about 0.52 μm), although precipitation of TiNbV-rich particles was also observed. Abbasi and Rainforth [40] studied the microstructure and mechanical properties in MoNbV microalloyed ferritic steel. Simultaneous additions of 0.08 wt. % Mo and 0.04 wt. % Nb to 0.12C-0.16V ferritic steel resulted in precipitation of MoNbVC and decreased size of VC particles, which was explained by the improved temperature stability and reduced coarsening rate of multi-element precipitates. Increased steel hardness with Nb and Mo microalloying was related to finer ferrite grain size and higher number density of VC particles in the NbMoV-microalloyed steel. In this work we advance the knowledge of multi-microalloyed steels in the following aspects: (i) compare the effects of minor Mo and Cr additions on phase transformation and particle precipitation in low carbon NbV-microalloyed bainitic steels; (ii) analyse the microstructure-property relationship in the newly developed steels with 700–850 MPa of yield stress; and (iii) investigate the effect of high temperature strain (in the recrystallization temperature region) on room temperature microstructure and mechanical properties. The effect of high temperature (>1000 ◦C) strain is important to study because increased strain values may enhance recrystallization of austenite (refine grain size) and accelerate precipitation of MoNbV-rich particles [10,23,28,41] (reduce Mo solid solute concentrations). Consequently, the grain size, precipitate number density and solute atom concentrations will affect the ambient temperature mechanical properties.

#### **2. Materials and Methods**

Two steels containing 0.08C, 1.5Mn, 0.3Si, 0.2Ni, 0.03Al, 0.003S, 0.015P, 0.01N, 0.06Nb, 0.12V and either 0.3Cr-0.2Mo or 0.5Cr-0Mo (wt. %), denoted below as MoNbV-steel and CrNbV-steel respectively, were melted in a 60 kg induction furnace and cast as 75 × 100 × 150 mm3 blocks by Hycast Metals Pty, Sydney, Australia. The blocks were homogenised at 1250 ◦C for 30 h, to equalise chemical composition, then forged in the temperature range of 1250–900 ◦C along the 100 mm side to 28 mm plate thickness, to assure 3.5 times reduction of the as-cast microstructure. The forged plates were cut into standard 20 × 15 × 10 mm3 Gleeble samples. Thermomechanical processing in Gleeble (manufactured by Dynamic Systems Inc., Poestenkill, NY, USA) was conducted using two schedules:

The processing schedule parameters (deformation temperature range, total strain and strain per pass, strain rate, and cooling rate between passes) have been defined to model the industrial rolling process within reasonable limits of the Gleeble simulator.

Microstructure characterisation for the four studied conditions was carried out using optical, scanning (SEM) and transmission (TEM) electron microscopy. For optical and SEM microscopy, the Gleeble samples were cut parallel to the normal direction (ND)–rolling direction (RD) plane, where ND is the compression direction and RD represents the rolling direction in Gleeble simulation. For TEM and tensile properties testing the samples were cut parallel to the normal direction (ND)–transverse direction (RD) plane. Optical and SEM sample preparation included polishing with SiC papers and diamond suspensions followed by etching with 5% Nital. Foils for TEM were prepared by hand polishing with a number of SiC papers, pre-thinning on a dimple grinder, and ion milling on a Gatan PIPS machine (manufactured by Gatan, Pleasanton, CA, USA). Optical microscopy was conducted on a Leica DM6000M microscope (manufactured by Leica Microsystems, Wetzlar, Germany) equipped with Leica Application Suite (LAS) 4.0.0 image processing software (developed by Leica Microsystems). Scanning electron microscopy was carried out using a JEOL 7001F FEG scanning electron microscope (manufactured by JEOL, Tokyo, Japan) operating at 15 kV for imaging and 7 kV for energy dispersive X-ray spectroscopy (EDS) of precipitates. For the determination of size of bainitic ferrite (the shortest distance between the martensitic grains) more than 400 randomly located areas were manually measured in SEM images for each of four studied conditions. In the SEM visible size range precipitation was scarce. Thus, only a limited number of 50 particles was analysed for the determination of precipitate sizes, number density and area fraction values for each of four studied conditions. The EDS semi-quantitative point analysis was carried out for 20 particles for each studied condition using an AZtec 2.0 Oxford SEM EDS system (manufactured by Oxford Instruments, Abingdon, UK). Transmission electron microscopy was conducted on a JEOL JEM2010 TEM microscope (manufactured by JEOL, Tokyo, Japan). For the analysis of <15 nm particle parameters, 200–500 precipitates were imaged for each of four studied conditions. The precipitates type was analyzed using selected area diffraction. The foil thickness was measured to be ~80 nm; a convergent beam diffraction technique was applied for this measurement [42]. Imaging of dislocation structure was performed for the beam direction being close to [011] grain zone axis. Tensile testing for the four studied conditions was carried out on a Kammrath and Weiss GmbH tensile stage. Testing was performed using 3 mm wide, 1 mm thick and 7 mm gauge length flat specimens. The constant crosshead speed of 7 μm·s−1 was applied and resulted in 1 × 10−3 s−1 strain rate. Two specimens were tested per condition.

#### **3. Results**

#### *3.1. Grain Structure and Phase Balance*

Optical, SEM and TEM microscopy showed in both steels a microstructure of mixed granular bainite and bainitic ferrite (Figure 1). Blocky or elongated martensite was present as the second phase. In both steels, the martensite crystals comprise a number of sub-grains with low angle (~10◦) boundaries between them in accordance with possible intervariant misorientation of 10.53◦ [43] for Kurdjumov–Sachs relationship between the parent austenite (face centred cubic (fcc) crystal structure) and product martensite (body centred cubic (bcc) crystal structure, observe the rotation of diffraction patterns in neighbouring sub-grains in Figures 2 and 3). The diffraction analysis (points A, B and C in Figure 2b, and points A and B in Figure 3b) confirmed the bcc type of crystal structure of martensite. Retained austenite was not observed. The average size of bainitic ferrite (the shortest distance between bainitic ferrite-martensite boundaries across the bainitic ferrite area) was measured to be below 1 μm (Table 1). The variation in the average sizes and shape of size distributions (Figure 4b) of bainitic ferrite with steel composition and processing schedule was insignificant and could result from the measurement error. However, a noticeable variation in the size of martensite grains was observed (Figure 4c,d). Thus, for low strain processing, the maximum size of blocky grains was 16% smaller and the maximum length of elongated grains was 40% shorter in the CrNbV-steel. For high strain processing an opposite trend was observed: the average and maximum sizes of blocky grains were 64% and 3.7 times, respectively, larger in the CrNbV-steel; and average and maximum length of elongated

grains were 20% and 30%, respectively, larger in the CrNbV-steel. In addition for high strain processing, the total fraction of martensite was 1.5 times higher in the CrNbV-steel.

 **Figure 1.** Optical images of microstructures in (**a**,**b**) MoNbV-steel and (**c**,**d**) CrNbV-steel after (**a**,**c**) low and (**b**,**d**) high strain processing.

 **Figure 2.** TEM (**a**) bright field image of microstructure and (**b**) dark field image of martensite in MoNbV-steel after low strain processing; (**b**) is from the white frame in (**a**); A, B and C diffraction patterns were taken from the corresponding points in the dark field image.

**Figure 3.** TEM (**a**) bright field image of microstructure and (**b**) dark field image of martensite in CrNbV-steel after low strain processing; (**b**) is from the white frame in (**a**); A and B diffraction patterns were taken from the corresponding points in the dark field image.

**Figure 4.** (**a**) A representative SEM image of microstructure in CrNbV-steel and size frequency distributions of (**b**) bainitic ferrite; (**c**) blocky martensite and (**d**) elongated martensite for four studied conditions.

**Table 1.** Microstructural parameters and mechanical properties of the studied steels.