**2.1.1 Chitosan to DNA complexation ratio, (N/P)c**

Stable DNA-chitosan polyplexes, ensuring a complete DNA compaction, are usually formed when chitosan is added in molar excess relative to the negatively charged DNA; however, excessive positive charges might lead to cytotoxicity since negatively charged cell membranes are prone to be damaged in the presence of cationic, extracellular compounds (Thomas & Klibanov, 2003). Consequently, finding a molar ratio exhibiting stable complexes at the lowest possible chitosan concentration becomes important. Such a concentration, presenting this mandatory characteristic, is what we define as the chitosan to DNA complexation ratio (N/P)c.

a. Determination of the (N/P)c via static light scattering (SLS)

386 Non-Viral Gene Therapy

It has to be noted that our characterizations were conducted as a function of two distinct cation-to-anion ratios reported in either molar (i.e., the N/P ratio, sections 2.1 and 2.2) or mass units (i.e., the L/D ratio, section 2.3). To get a complete description of the sample preparation, experimental procedures, and data analysis of the results here exposed, the reader is encouraged to consult our published papers (Alatorre-Meda et al., 2009; Alatorre-Meda et al., 2010a, 2010b; Alatorre-Meda et al., 2011). Outstanding results are

Upon mixing, oppositely charged polyelectrolytes interact electrostatically and form complexes in a process that is promoted by an increase in entropy which is due to a release of counterions (Manning, 1978; Matulis et al., 2000). Accordingly, polycation molecular parameters such as charge density and valence have gained attention in recent reports

The role of chitosan charge density is well established. It is accepted that the high charge density of chitosan at pHs below its pKa results beneficial for polyplex preparation, and also that its low charge density at pH 7.4 contributes to a low polyplex cytotoxicity and facilitates the intracellular release of DNA from the complex after its endocytotic cellular uptake (Strand et al., 2010). By contrast, the role of chitosan valence on transfection efficiency is contradictory. Namely, while several studies promote the use of high Mw chitosans (Huang et al., 2005; MacLaughlin et al., 1998), some other publications report that lower Mw chitosans are superior for gene transfer (Koping-Hoggard et al., 2003; Lavertu et al.,

Aiming to draw general conclusions about the feasibility of using chitosan as a gene carrier, we characterized the DNA complexation and transfection mediated by three chitosans presenting different Mw (three different valences) at three different pHs of 5.0, 6.0, and 6.5 (three different charge densities). Table 1 summarizes the physical characteristics of the

CHITOSAN [η] (dl g-1) Mw (kDa) Label Low viscous 4.42 + 0.01 111 + 2 C(689) Middle viscous 7.85 + 0.24 266 + 14 C(1652) Highly viscous 11.40 + 0.24 467 + 18 C(2901)

Stable DNA-chitosan polyplexes, ensuring a complete DNA compaction, are usually formed when chitosan is added in molar excess relative to the negatively charged DNA; however, excessive positive charges might lead to cytotoxicity since negatively charged cell membranes are prone to be damaged in the presence of cationic, extracellular compounds (Thomas & Klibanov, 2003). Consequently, finding a molar ratio exhibiting stable complexes at the lowest possible chitosan concentration becomes important. Such a concentration, presenting this mandatory characteristic, is what we define as the chitosan to DNA

presented below.

2006).

chitosans employed.

Table 1. Chitosans employed.

complexation ratio (N/P)c.

**2.1.1 Chitosan to DNA complexation ratio, (N/P)c** 

**2.1 The DNA-chitosan system** 

(Danielsen et al., 2004; Maurstad et al., 2007).

It is well accepted that linear, highly charged polyelectrolytes, at the dilute and semi-dilute regimes, can interact via a Coulombic repulsive potential which is strong enough to keep the polymer chains elongated and widely separated, although these interactions be partly screened by non-condensed counterions in solution (Manning, 1978). Polyelectrolytes in such concentration regimes produce in consequence very small scattering signals when irradiated with any source of light (Drifford & Dalbiez, 1984). The situation is rather distinct when polyelectrolytes interact one with each other or are complexed with external agents; in such a case they scatter higher amounts of light when irradiated (Drifford & Dalbiez, 1984). Based on these foundations we analyzed our polyplexes via SLS in order to follow the complexation process. SLS has proved to be a suitable tool to detect structural changes in linear biopolymers expected to be compacted provided that upon structural changes (such as the coil-globule transition observed during DNA compaction) they must scatter light to different extents.

To determine (N/P)c, we tested numerous chitosan/DNA formulations with varying N/P ratios (at a constant DNA concentration) in terms of the chitosan charge density and valence. What we found by SLS was that there exist, indeed, a molar ratio from which the structural conformation of the polyplexes remains apparently constant independently of further addition of chitosan (i.e., presenting non-accentuated changes in light scattering intensity). That molar ratio, defined as (N/P)c, proved to be strongly influenced by the chitosan charge density adopting values of around 1.5, 2.0, and 6.0 for the pHs of 5.0, 6.0, and 6.5, respectively. Our estimations, inferred from figure 1, can be discussed as follows.

Figure 1 presents the light scattering intensity of the DNA-C(1652) polyplexes as a function of the N/P ratio for the three studied pHs. This figure reflects various features worth analyzing. Interestingly, the system revealed plots similar in shape, but different in I values. The plots collected at pH 5.0 and 6.0 have I values one close to the other, whereas the plot at pH 6.5 has higher I values over the whole range of N/P studied. In all three plots three distinct regions can be identified, namely at N/P = 0, at 0 < N/P ≲ 2, and at N/P >2. At N/P = 0, the system containing pure DNA shows the intensity at least five times lower than the samples at N/P > 2, indicating no aggregation. However, upon addition of chitosan to DNA, the intensity increases sharply with the maximum at N/P around 1-2 to finally level off at N/P > 2. This peculiar and interesting behavior observed when chitosan concentration in the system relative to DNA is around 1-2, is suggestive of the formation of some kind of complex structures between DNA and chitosan at this region, large in size, possibly aggregates that are responsible for the dispersion of higher amounts of light. Finally at N/P > 2, the intensity I reaches a constant value revealing the presence of well-formed, stable DNA-chitosan polyplexes with regular sizes. The N/P ratio marking the onset of the constant value in I is denoted as the (N/P)c. Very importantly, these results suggest that as the pH of the medium increases larger amounts of chitosans are required to completely compact the given amount of DNA. This phenomenon can be explained by the fact that at pHs close to its pKa (6.3–6.5), chitosan undergoes a decrease in its charge density due to the neutralization of its amino groups (Kumar et al., 2004), a feature that becomes even more pronounced for chitosans with higher molecular weights (higher valences) (MacLaughlin et al., 1998). Therefore, and having in mind that one of the driving forces of polyelectrolyte complexation is the release of counterions from the polyanion–polycation pair (Manning, 1978; Matulis et al., 2000), it is not strange that the binding affinity between DNA and chitosan lowered as the pH increased and got close to 6.5. Likewise, the highest intensities

Polycation-Mediated Gene Delivery: The Physicochemical Aspects Governing the Process 389

concentration of 0.104 mM, corresponding to the N/P ratio of 0.82. A similar inflection in a conductivity plot during DNA compaction upon addition of a cationic vector has been observed elsewhere (Rodriguez-Pulido et al., 2008). The authors suggested that the increase in conductivity related to the counterion release from the polycation injected is accentuated by the release of counterions resulting from the complexation process (in our case Na+ from DNA and CH3COO− from chitosan) thereby justifying a higher slope in the conductivity plot below the inflection point. On the other hand, once the inflection occurred, the lower slope can be attributed to the fact that only the counterions coming from the chitosan dissociation now contribute to the conductivity of the solution. This change in slope of the conductivity plot can in consequence be considered as the point from which DNA is

Compared to the DNA compaction ratio we determined by SLS ((N/P)c ~ 1.5) , the lower ratio of N/P = 0.82 here depicted can be ascribed to the difference in ionic strength of the media used in both experiments and to the fact that contrary to SLS, in the conductometry experiment the complex formation was run at constant stirring.1 The other two chitosans, C(689) and C(1652), although with slight differences in the conductivity values, revealed the inflection point at exactly the same N/P ratio as compared to C(2901) (plots not shown).

**0.00 0.05 0.10 0.15 0.20 0.25 0.30 0.35 0.40**

**C(2901), mM**

Reproduced from (Alatorre-Meda et al., 2011) with permission of Elsevier BV in the format Journal via

Fig. 2. Electrical conductivity, κ, vs. C(2901) concentration. Filled and empty squares stand for the addition of chitosan to a DNA and to a pure buffer solution, respectively. (Note the

<sup>1</sup> Sample preparation for conductometry experiments is described in (Alatorre-Meda et al., 2011). To

consult the experimental conditions for SLS go to (Alatorre-Meda et al., 2009).

κ**POLYPLEX,** μ**S cm-1**

compacted, namely (N/P)c (Rodriguez-Pulido et al., 2008).

*(N/P)c* **= 0.82**

κ**CHITOSAN,** μ**S cm-1**

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difference in scales).

observed for the system at pH 6.5 over the whole range of N/P can be attributed to a lower chitosan solubility (also ascribed to pHs close to the pKa), a fact that leads to the formation of polyplexes of sizes larger than those at lower pHs, as described below (MacLaughlin et al., 1998; Mumper al., 1995). The DNA-C(689) and -C(2901) systems revealed the same behavior (data not included).

Adapted from (Alatorre-Meda et al., 2009).

Fig. 1. I vs. N/P of the DNA-C(1652) polyplexes at pH = 5.0 (squares), pH = 6.0 (circles), and pH = 6.5 (triangles).

#### b. Determination of the (N/P)c by conductometry

Electrostatic interactions between oppositely charged polyelectrolytes entail a release of counterions (Manning, 1978; Matulis et al., 2000). The tracking of this release by means of conductometry can be employed as a tool to characterize the DNA compaction process (Rodriguez-Pulido et al., 2008). To confirm the (N/P)c values obtained by SLS we measured the change in conductivity provoked by the addition of chitosan solutions to both, DNA and pure buffer solutions. Compared to those of SLS, the conductometry results, depicted in figure 2, reveal (N/P)c values slightly lower. Main findings as well as a possible explanation to the observed differences are exposed below.

Figure 2 presents a representative plot of the electrical conductivity, κ, as a function of the polycation concentration, recorded for C(2901) at pH 5.0. The filled and empty squares stand for the DNA and pure buffer reservoir solutions, respectively. As can be seen from this figure, in the buffer solution alone the conductivity increased linearly with the chitosan concentration (empty squares) indicating that no aggregation took place under the whole range of the polycation concentration. In the DNA solution (filled squares) by contrast, the conductivity grew linearly, however, with a clear change in slope at the chitosan

observed for the system at pH 6.5 over the whole range of N/P can be attributed to a lower chitosan solubility (also ascribed to pHs close to the pKa), a fact that leads to the formation of polyplexes of sizes larger than those at lower pHs, as described below (MacLaughlin et al., 1998; Mumper al., 1995). The DNA-C(689) and -C(2901) systems revealed the same

**0 5 10 15 20**

*N/P*

Fig. 1. I vs. N/P of the DNA-C(1652) polyplexes at pH = 5.0 (squares), pH = 6.0 (circles), and

Electrostatic interactions between oppositely charged polyelectrolytes entail a release of counterions (Manning, 1978; Matulis et al., 2000). The tracking of this release by means of conductometry can be employed as a tool to characterize the DNA compaction process (Rodriguez-Pulido et al., 2008). To confirm the (N/P)c values obtained by SLS we measured the change in conductivity provoked by the addition of chitosan solutions to both, DNA and pure buffer solutions. Compared to those of SLS, the conductometry results, depicted in figure 2, reveal (N/P)c values slightly lower. Main findings as well as a possible explanation

Figure 2 presents a representative plot of the electrical conductivity, κ, as a function of the polycation concentration, recorded for C(2901) at pH 5.0. The filled and empty squares stand for the DNA and pure buffer reservoir solutions, respectively. As can be seen from this figure, in the buffer solution alone the conductivity increased linearly with the chitosan concentration (empty squares) indicating that no aggregation took place under the whole range of the polycation concentration. In the DNA solution (filled squares) by contrast, the conductivity grew linearly, however, with a clear change in slope at the chitosan

behavior (data not included).

*I***, kC s-1**

Adapted from (Alatorre-Meda et al., 2009).

b. Determination of the (N/P)c by conductometry

to the observed differences are exposed below.

pH = 6.5 (triangles).

concentration of 0.104 mM, corresponding to the N/P ratio of 0.82. A similar inflection in a conductivity plot during DNA compaction upon addition of a cationic vector has been observed elsewhere (Rodriguez-Pulido et al., 2008). The authors suggested that the increase in conductivity related to the counterion release from the polycation injected is accentuated by the release of counterions resulting from the complexation process (in our case Na+ from DNA and CH3COO− from chitosan) thereby justifying a higher slope in the conductivity plot below the inflection point. On the other hand, once the inflection occurred, the lower slope can be attributed to the fact that only the counterions coming from the chitosan dissociation now contribute to the conductivity of the solution. This change in slope of the conductivity plot can in consequence be considered as the point from which DNA is compacted, namely (N/P)c (Rodriguez-Pulido et al., 2008).

Compared to the DNA compaction ratio we determined by SLS ((N/P)c ~ 1.5) , the lower ratio of N/P = 0.82 here depicted can be ascribed to the difference in ionic strength of the media used in both experiments and to the fact that contrary to SLS, in the conductometry experiment the complex formation was run at constant stirring.1 The other two chitosans, C(689) and C(1652), although with slight differences in the conductivity values, revealed the inflection point at exactly the same N/P ratio as compared to C(2901) (plots not shown).

Reproduced from (Alatorre-Meda et al., 2011) with permission of Elsevier BV in the format Journal via Copyright Clearance Center.

Fig. 2. Electrical conductivity, κ, vs. C(2901) concentration. Filled and empty squares stand for the addition of chitosan to a DNA and to a pure buffer solution, respectively. (Note the difference in scales).

<sup>1</sup> Sample preparation for conductometry experiments is described in (Alatorre-Meda et al., 2011). To consult the experimental conditions for SLS go to (Alatorre-Meda et al., 2009).

Polycation-Mediated Gene Delivery: The Physicochemical Aspects Governing the Process 391

**100 150 200 250 300 350 400 450 500**

*Mw***, kDa**

Reproduced from (Alatorre-Meda et al., 2009) with permission of Elsevier BV in the format Journal via

**0 4 8 12 16 20**

Reproduced from (Alatorre-Meda et al., 2011) with permission of Elsevier BV in the format Journal via

Fig. 4. ζ-potential of DNA-chitosan polyplexes vs. N/P. DNA–C(689) (squares), DNA– C(1652) (circles), and DNA–C(2901) (triangles) are plotted. The dotted line stands for the

*N/P*

**Mean = 16 Std dev = 1.62**

**DNA**

**-60**

DNA solution (ζ-potential = -55 mV).

**0**

**4**

ζ

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 **potential, mV**

**8**

**12**

**16**

**20**

Fig. 3. RH of DNA-chitosan polyplexes, vs. Mw of chitosan.

*R*

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*H***, nm**

#### **2.1.2 Time stability and size**

To determine the time stability of the polyplexes, we measured by dynamic light scattering (DLS) the hydrodynamic radius, RH, of a sample presenting an N/P > (N/P)c (more specifically, N/P = 6) during a period of 6 days at the three different pHs of interest. In general, the polyplexes, regardless of charge density and valence, presented constant sizes with fluctuations lower than a 10% (data not shown). Thus, and provided that small fluctuations as those observed in our measurements are most likely related to the nature of the DLS technique, the polyplexes can be considered as stable with time.

Considering their time stability, to calculate the characteristic size of the polyplexes we simply averaged the RH values obtained along the testing time. The polyplex sizes thereby obtained were in the range of 187 + 21 < RH < 246 + 13 nm, in good agreement with results previously reported (Mumper et al., 1995). However, contrary to what we initially expected, the size of the polyplexes was found to be dependent on the chitosan valence, following a linear trend with the chitosan molecular weight.2 Figure 3 depicts the average size of the polyplexes (regardless of chitosan charge density) as a function of the chitosan molecular weight.

Increasing trends in the complex size with polycation Mw, such as that observed in figure 3, are well documented in the literature. For the case of chitosan, it has been demonstrated that upon increasing in its chain length, the influence of the charge density with its correspondent entropy gain decrease. It is likely that the restriction of the polycation chain upon complexation becomes more important, giving rise to a different complexation behavior of high Mw chitosans compared to low Mw chitosans (Danielsen et al., 2004; Maurstad et al., 2007). Furthermore, the intuitive assumption that a higher Mw chitosan can interact better with DNA (due to its expected higher valence), and thus condense it more efficiently than a chitosan of a lower Mw is outweighed by the fact that a higher molecular weight chitosan is less soluble, and as a result, an increase in complex diameter or even complex aggregation may result (MacLaughlin et al., 1998; Mumper et al., 1995).

A general conclusion drawn throughout these sections is that an increased charge density of chitosan, resulting from a lowering of pH, leads to a greater binding affinity between chitosan and DNA as fewer chitosan is required to reach the complexation and the complexes thereby formed are more stable (Alatorre-Meda et al., 2009). Consequently, and in order to further understand the general aspects involved in the DNA–chitosan interactions, in subsequent studies we focused on working at acidic conditions exclusively (Alatorre-Meda et al., 2011). The influence of chitosan valence at those conditions on complex physicochemical properties other than size is described below.

#### **2.1.3 Surface charge**

For many polyplexes the cross-over from a negative to a positive ζ-potential occurs at or very close to the isoneutrality point (N/P)ϕ. (N/P)ϕ is defined as the point at which the N/P ratio of the polyplex equals 1, that is, the ratio where the negative charges of DNA are stoichiometrically neutralized by the positive charges of the polycation (De Smedt et al., 2000).

<sup>2</sup> In general, it is believed that polycations with higher charge densities and valences should produce smaller DNA complexes.

To determine the time stability of the polyplexes, we measured by dynamic light scattering (DLS) the hydrodynamic radius, RH, of a sample presenting an N/P > (N/P)c (more specifically, N/P = 6) during a period of 6 days at the three different pHs of interest. In general, the polyplexes, regardless of charge density and valence, presented constant sizes with fluctuations lower than a 10% (data not shown). Thus, and provided that small fluctuations as those observed in our measurements are most likely related to the nature of

Considering their time stability, to calculate the characteristic size of the polyplexes we simply averaged the RH values obtained along the testing time. The polyplex sizes thereby obtained were in the range of 187 + 21 < RH < 246 + 13 nm, in good agreement with results previously reported (Mumper et al., 1995). However, contrary to what we initially expected, the size of the polyplexes was found to be dependent on the chitosan valence, following a linear trend with the chitosan molecular weight.2 Figure 3 depicts the average size of the polyplexes (regardless of chitosan charge density) as a function of the chitosan

Increasing trends in the complex size with polycation Mw, such as that observed in figure 3, are well documented in the literature. For the case of chitosan, it has been demonstrated that upon increasing in its chain length, the influence of the charge density with its correspondent entropy gain decrease. It is likely that the restriction of the polycation chain upon complexation becomes more important, giving rise to a different complexation behavior of high Mw chitosans compared to low Mw chitosans (Danielsen et al., 2004; Maurstad et al., 2007). Furthermore, the intuitive assumption that a higher Mw chitosan can interact better with DNA (due to its expected higher valence), and thus condense it more efficiently than a chitosan of a lower Mw is outweighed by the fact that a higher molecular weight chitosan is less soluble, and as a result, an increase in complex diameter or even complex aggregation may result (MacLaughlin et al., 1998; Mumper

A general conclusion drawn throughout these sections is that an increased charge density of chitosan, resulting from a lowering of pH, leads to a greater binding affinity between chitosan and DNA as fewer chitosan is required to reach the complexation and the complexes thereby formed are more stable (Alatorre-Meda et al., 2009). Consequently, and in order to further understand the general aspects involved in the DNA–chitosan interactions, in subsequent studies we focused on working at acidic conditions exclusively (Alatorre-Meda et al., 2011). The influence of chitosan valence at those conditions on

For many polyplexes the cross-over from a negative to a positive ζ-potential occurs at or very close to the isoneutrality point (N/P)ϕ. (N/P)ϕ is defined as the point at which the N/P ratio of the polyplex equals 1, that is, the ratio where the negative charges of DNA are stoichiometrically neutralized by the positive charges of the polycation (De Smedt et

2 In general, it is believed that polycations with higher charge densities and valences should produce

complex physicochemical properties other than size is described below.

the DLS technique, the polyplexes can be considered as stable with time.

**2.1.2 Time stability and size**

molecular weight.

et al., 1995).

**2.1.3 Surface charge** 

smaller DNA complexes.

al., 2000).

Reproduced from (Alatorre-Meda et al., 2009) with permission of Elsevier BV in the format Journal via Copyright Clearance Center.

Fig. 3. RH of DNA-chitosan polyplexes, vs. Mw of chitosan.

Reproduced from (Alatorre-Meda et al., 2011) with permission of Elsevier BV in the format Journal via Copyright Clearance Center.

Fig. 4. ζ-potential of DNA-chitosan polyplexes vs. N/P. DNA–C(689) (squares), DNA– C(1652) (circles), and DNA–C(2901) (triangles) are plotted. The dotted line stands for the DNA solution (ζ-potential = -55 mV).

Polycation-Mediated Gene Delivery: The Physicochemical Aspects Governing the Process 393

It has been reported that the DNA complexation with chitosans might result in a blend of structures: toroids, rods, and globules, with the relative amounts of the different structures apparently depending on the actual chitosan, the charge ratio, and solution properties like pH and ionic strength (Danielsen et al., 2004; Maurstad et al. 2007). What we observe from figure 5A and B is a heterogeneous population of polyplexes with particle sizes ranging from 250 to 500 nm in good agreement with the DLS results (see section 2.1.2). Both images depict polyplexes with a brush-like conformation where globules/aggregates comprise a dense core that is surrounded by a "hairy" shell of polymer chains. This globular conformation has been reported as characteristic of the DNA complexation with high molecular weight chitosans (Mw> 100 kDa) (Danielsen et al., 2004; Maurstad et al., 2007); by contrast, complexes formed with lower molecular weight chitosans adopt toroid- and rodlike conformations (Maurstad et al., 2007). Similar brush-like structures were also obtained for the DNA complexation with transferrin-poly(l-lysine) conjugates. In this case, the complex morphology was found to depend on the conjugate to DNA ratio (Wagner et al., 1991). Carnerup and coworkers suggest that the significant morphological rearrangement undergone by DNA when it is condensed with low Mw polycations is because of the low charge density of the polycation in question (Carnerup et al., 2009). For toroidal aggregates to form, the electrostatic attraction has to be moderate; that is, a balance between mobility and high binding affinity of DNA to the polymer has to exist. In such a system, the condensed DNA chains will be able to arrange into a toroid. On the contrary, if the charge density of the polymer is too high (as expected for chitosan at pH 5), the DNA chains will entangle with the polymer ones, forming globular aggregates (Carnerup et al., 2009). Therefore, provided that our polyplexes proved to be stable (see section 2.1.2), the "not so tight" DNA complexation they present should be beneficial for DNA transfection (Tros de

Very importantly, the morphological structure of our system depicted by TEM and AFM in conjunction with the markedly positive ζ-potentials obtained for the polyplexes at this high N/P ratio (see the previous section) appear to be in line with the core–shell structure proposed for polycation-excessive DNA complexes (A. V. Kabanov & V. A. Kabanov, 1998). This model states that DNA is condensed in the inner part of the polyplex by the binding of short segments of a large number of polycation chains, whereas the remaining segments of these same chains are expected to be free in the outer part of the polyplexes giving rise to

Once the complex is released from the endosome, DNA must disassemble from its vector to be accessible to the cell machinery responsible for translating the enclosed information. For the case of polyplexes, the DNA decompaction (disassembly) is a process substantially dependent on i) the DNA-vector binding affinity, ii) the complexation thermodynamics, and iii) the solution conditions (Carlstedt et al., 2010; Prevette et al., 2007). Provided that acidic conditions demonstrated to be optimal for polyplex formation (see previous sections), we evaluated the DNA-chitosan binding affinity and complexation thermodynamics at a solution pH of 5.0. As depicted by isothermal titration calorimetry (ITC), lower valence chitosans demonstrated to have a higher binding affinity for DNA. Main results are

markedly positive polyplex surface charges (A. V. Kabanov & V. A. Kabanov, 1998).

**2.1.5 Binding affinity and complexation thermodynamics**

Ilarduya et al., 2010).

described below.

In the present study, the characterization was done in the range 1 ≤ N/P ≤ 20 for all polyplexes. The ζ-potential of the polyplexes is plotted as a function of N/P in figure 4. In general, all polyplexes presented a positive, stable ζ-potential from N/P ratios as low as (N/P)c confirming that DNA is completely compacted independently of further addition of chitosan. Main findings can be discussed as follows.

It can be seen from figure 4 that at N/P = 1 all polyplexes, in particular those formed with C(1652) and C(2901), reveal a lower ζ-potential as compared to the rest of compositions. The cationic vector-mediated DNA coil to globule transition demonstrated by other authors (Dias et al., 2005) in conjunction with the base line-absent DLS correlation functions we obtained for these systems at ratios N/P ≤ 1 (Alatorre-Meda et al., 2009), may provide an explanation to this feature. Apparently, larger amounts of chitosan are needed to completely compact the DNA and in consequence populations entailing varying extents of DNA compaction are expected to be present in the bulk. On the other hand, at ratios higher than (N/P)ϕ, all polyplexes reach a plateau around 16mV regardless of chitosan Mw, which is in good agreement with other DNA-polycation systems (Tang & Szoka, 1997). This positive ζpotential of the polyplexes suggests that the DNA compaction is completely achieved with chitosan chains probably pointing to the outer part of the polyplexes as inferred by other authors (Koping-Hoggard et al., 2003).

#### **2.1.4 Structural organization**

Imaging techniques can detect, localize, and analyze individual aggregates of a heterogeneous population, thereby revealing events that would otherwise be hidden. In this context transmission electron- and atomic force microscopy (TEM and AFM) are frequently used in parallel for the visual characterization of biological molecules (Arakawa et al., 1992; Lin & Goh, 2002). Figure 5 presents typical TEM (A) and AFM (B) images obtained for the DNA–C(689) polyplexes at N/P = 20. This figure reflects that the polyplexes adopt a peculiar brush-like conformation in which DNA is apparently confined to the interior of the complex although not fully compacted. The reason why of this polyplex conformation as well as the implications it might have on transfection are discussed below.

Reproduced from (Alatorre-Meda et al., 2011) with permission of Elsevier BV in the format Journal via Copyright Clearance Center.

Fig. 5. TEM (A) and height AFM (B) images of DNA-C(689) polyplexes, N/P = 20. The bar next to (B) represents the Z scale in nm.

In the present study, the characterization was done in the range 1 ≤ N/P ≤ 20 for all polyplexes. The ζ-potential of the polyplexes is plotted as a function of N/P in figure 4. In general, all polyplexes presented a positive, stable ζ-potential from N/P ratios as low as (N/P)c confirming that DNA is completely compacted independently of further addition of

It can be seen from figure 4 that at N/P = 1 all polyplexes, in particular those formed with C(1652) and C(2901), reveal a lower ζ-potential as compared to the rest of compositions. The cationic vector-mediated DNA coil to globule transition demonstrated by other authors (Dias et al., 2005) in conjunction with the base line-absent DLS correlation functions we obtained for these systems at ratios N/P ≤ 1 (Alatorre-Meda et al., 2009), may provide an explanation to this feature. Apparently, larger amounts of chitosan are needed to completely compact the DNA and in consequence populations entailing varying extents of DNA compaction are expected to be present in the bulk. On the other hand, at ratios higher than (N/P)ϕ, all polyplexes reach a plateau around 16mV regardless of chitosan Mw, which is in good agreement with other DNA-polycation systems (Tang & Szoka, 1997). This positive ζpotential of the polyplexes suggests that the DNA compaction is completely achieved with chitosan chains probably pointing to the outer part of the polyplexes as inferred by other

Imaging techniques can detect, localize, and analyze individual aggregates of a heterogeneous population, thereby revealing events that would otherwise be hidden. In this context transmission electron- and atomic force microscopy (TEM and AFM) are frequently used in parallel for the visual characterization of biological molecules (Arakawa et al., 1992; Lin & Goh, 2002). Figure 5 presents typical TEM (A) and AFM (B) images obtained for the DNA–C(689) polyplexes at N/P = 20. This figure reflects that the polyplexes adopt a peculiar brush-like conformation in which DNA is apparently confined to the interior of the complex although not fully compacted. The reason why of this polyplex conformation as

Reproduced from (Alatorre-Meda et al., 2011) with permission of Elsevier BV in the format Journal via

Fig. 5. TEM (A) and height AFM (B) images of DNA-C(689) polyplexes, N/P = 20. The bar

well as the implications it might have on transfection are discussed below.

chitosan. Main findings can be discussed as follows.

authors (Koping-Hoggard et al., 2003).

**2.1.4 Structural organization**

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next to (B) represents the Z scale in nm.

It has been reported that the DNA complexation with chitosans might result in a blend of structures: toroids, rods, and globules, with the relative amounts of the different structures apparently depending on the actual chitosan, the charge ratio, and solution properties like pH and ionic strength (Danielsen et al., 2004; Maurstad et al. 2007). What we observe from figure 5A and B is a heterogeneous population of polyplexes with particle sizes ranging from 250 to 500 nm in good agreement with the DLS results (see section 2.1.2). Both images depict polyplexes with a brush-like conformation where globules/aggregates comprise a dense core that is surrounded by a "hairy" shell of polymer chains. This globular conformation has been reported as characteristic of the DNA complexation with high molecular weight chitosans (Mw> 100 kDa) (Danielsen et al., 2004; Maurstad et al., 2007); by contrast, complexes formed with lower molecular weight chitosans adopt toroid- and rodlike conformations (Maurstad et al., 2007). Similar brush-like structures were also obtained for the DNA complexation with transferrin-poly(l-lysine) conjugates. In this case, the complex morphology was found to depend on the conjugate to DNA ratio (Wagner et al., 1991). Carnerup and coworkers suggest that the significant morphological rearrangement undergone by DNA when it is condensed with low Mw polycations is because of the low charge density of the polycation in question (Carnerup et al., 2009). For toroidal aggregates to form, the electrostatic attraction has to be moderate; that is, a balance between mobility and high binding affinity of DNA to the polymer has to exist. In such a system, the condensed DNA chains will be able to arrange into a toroid. On the contrary, if the charge density of the polymer is too high (as expected for chitosan at pH 5), the DNA chains will entangle with the polymer ones, forming globular aggregates (Carnerup et al., 2009). Therefore, provided that our polyplexes proved to be stable (see section 2.1.2), the "not so tight" DNA complexation they present should be beneficial for DNA transfection (Tros de Ilarduya et al., 2010).

Very importantly, the morphological structure of our system depicted by TEM and AFM in conjunction with the markedly positive ζ-potentials obtained for the polyplexes at this high N/P ratio (see the previous section) appear to be in line with the core–shell structure proposed for polycation-excessive DNA complexes (A. V. Kabanov & V. A. Kabanov, 1998). This model states that DNA is condensed in the inner part of the polyplex by the binding of short segments of a large number of polycation chains, whereas the remaining segments of these same chains are expected to be free in the outer part of the polyplexes giving rise to markedly positive polyplex surface charges (A. V. Kabanov & V. A. Kabanov, 1998).
