DNA-based drug carriers and dynamic proteoids with tunable properties Liu, Yun
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Chapter 6
Dynamic Proteoids: Biodynamers Generated from Amino Acid-
or Dipeptide-Based Monomers
In this chapter, we designed and generated a range of doubly dynamic proteoid biodynamers based on the polycondensation of different types of amino acid and dipeptide hydrazides with a nonbiological aromatic dialdehyde and a biological aliphatic dialdehyde through formation of two types of reversible C=N bonds (imine and acylhrazone). By using the biological dialdehyde, biocompatibility of biodynamers formed was enhanced. Meanwhile, we compared the respective importance of the three factors that influence the polymerization and structure of the resulting biodynamer, including aromaticity, charge and the presence of hydroxyl group in the side chain of the amino acid.
Part of chapter will be submitted for publication:
6.1 Introduction
In chapter 5, we prepared dynamic proteoids through reversible polycondensation of a water-soluble amphiphilic dialdehyde with various bifunctional amino acid hydrazides.[1] We found that polymerization is driven by the self-organization/folding of the dynamic proteoids formed through hydrophobic interactions.[1–2] More importantly, side chains of amino acid hydrazides have a strong influence on the rates of polymerization, structures and dynamic properties of the resulting dynamic proteoids, including the aromaticity, charges and polarity.
Proteins consist, however, of amino acids with different side chains. Hence, it is necessary to evaluate their influence on polymerization and structure of the resulting dynamic proteoids, including aromaticity, charge and polarity of the amino acids. On the other hand, as the use of the nonbiological dialdehyde 1 leads to a decrease in biocompatibility of the resulting biodynamers, its replacement with a bio-based dialdehyde may circumvent this problem. Based on these considerations, we report here the design and synthesis of a series of novel dynamic proteoids through polycondensation of two dialdehydes (dialdehyde 1 is nonbiological and dialdehyde 2 is bio-derived) with various types of amino acid and dipeptide hydrazides bearing different side chains (Scheme 1).
6.2 Results and discussion
6.2.1 Molecular design of monomersThe formation of reversible C=N bonds, including both imine and acylhydrazone bonds, have been employed to generate dynamic proteoids. Moreover, the presence of two types of C=N bonds (true imines and acylhydrazones)[3] affords biodynamers of double dynamicity and pH-responsiveness and potentially a third form of dynamic behavior through structure-formation processes (conformational dynamics). Through the polycondensation of dialdehydes 1 or 2 with various amino acid or dipeptide hydrazides a–j (Scheme 1), dynamic proteoids with different structures and stabilities are formed in aqueous media under mildly acidic conditions (pD~5).
The aromatic nonbiological dialdehyde 1 consists of a tricyclic aromatic core and a hexaglyme chain. We have reported that polycondensation of dialdehyde 1 with amino acid hydrazides is driven by hydrophobic interactions deriving from the tricyclic core while the hexaglyme chains endows the biodynamers obtained with water-solubility and stabilizes them in aqueous media. Replacement of dialdehyde 1 with a furanose-based dialdehyde 2 should improve the biocompatibility of the resulting biodynamers. Moreover, hydroxyl groups in dialdehyde 2[4] can stabilize the
105 biodynamers through hydrogen bonds and/or OH−π interactions. Additionally, by comparing the aromatic dialdehyde 1 with the aliphatic dialdehyde 2, we will study the influence of the aromatic backbone on biodynamer formation.
N H O H O O 6 N H NH2 H2N O N HN N H NH2 H2N O N H NH2 H2N O ONa NH NH2 H2N O OH NH2 O 1 2 a b g f e d c H2N H N N H O O N NH HN N H2N H N N H O O NH2 NH2 H2N H N N H O O OH HO h H2N H N N H O O HN N OH H2N H N N H O O HN NH2 H2N H N N H O O OH NH2 j i O OH HO OH OH HO HO NH2 NH2 NH2 NH2 NH2 NH2
Scheme 1. Structures of dialdehydes 1 and 2, amino acid (a–d) and dipeptide hydrazides (e–f).
Our previous research demonstrates that three types of factors, including aromaticity, positive charge and hydroxyl groups, dramatically facilitate polymerization and stabilize structures of biodynamers through non-covalent interactions between side chains of amino acid hydrazides and dialdehyde 1, including π−π-stacking, cation–π interactions and hydrogen bonds. Furthermore, electrostatic forces dominate the polymerization when two oppositely charged amino acids are used.[1] Along these lines, we designed three categories of complementary monomers for the dialdehydes (Scheme 1): (1) amino acid hydrazides bearing an aromatic ring (a), a positive charge (b), a negative charge (c) and a hydroxyl group (d); (2) dipeptide hydrazides containing two aromatic rings (e), two negative charges (f) or two hydroxyl groups (g), which should represent an enhancement of one beneficial factor; (3) dipeptide hydrazides consisting of two different types of side chains, which
is a combination of two beneficial factors, such as an aromatic ring with a positive charge (h), an aromatic ring with a hydroxyl group (i), and a positive charge with a hydroxyl group (j). Polycondensation of these monomers with dialdehydes afforded a series of biodynamers with different structures. By studying the polymerization through 1H-NMR and characterizing the biodynamers generated via light scattering (LS) and cryo-transmission-electron microscopy (cyro-TEM), we evaluated the respective importance of the various factors on polymerization and structure of the resulting biodynamers, which could set the stage for the rational design of well-ordered nano-structures as novel biomaterials.
N H O H O O 6 H2N H N N H O O R2 R1 NH2 N N N O 6 H N O R2 N H N O R1 n + pD = 5
1 e-j poly(1-e) to poly(1-j)
Scheme 2. Schematic representation of the preparation of dynamic proteoids through reversible polycondensation of dialdehyde 1 with dipeptide hydrazides a–j.
Table 1. Consumption of dialdehydes 1 or 2 obtained by monitoring the signals from the aldehyde group by 1H-NMR spectroscopy (the integration of the signals from the aldehyde group at 5 min is set to 100%).
Samples Consumption of dialdehyde 1 (%) Samples Consumption of dialdehyde 1 (%)
poly(1-e) 100 poly(1-h) 79
poly(1-f) 50 poly(1-i) 97
poly(1-g) 95 poly(1-j) 64
6.2.2 Generation and characterization of biodynamers containing dialdehyde 1 We previously found the mechanism of polymerization to be nucleation-elongation (N-E), characterized by the formation of a critical size of polymer chain (nucleus) and elongation of the existing polymer, which is more favorable than initiation of a new chain. At pD 5, acylhydrazone formation proceeds readily and goes to completion, whereas the amine does not afford the corresponding imine. However, due to the hydrophobic and π−π-stacking interactions from the main chain, polymerization took place and generated thermodynamic biodynamers with well-ordered nano-structures. We also observed that aromaticity, positive charge and hydroxyl groups of the side chains of amino acidhydrazides significantly speed up polymerization and facilitate formation of the corresponding biodynamers through non-covalent interactions. To evaluate the respective importance of aromaticity, positive charge and hydroxyl group
107 in the formation of dynamic proteoids, biodynamers were synthesized through the polycondensation of dialdehyde 1 with dipeptide hydrazides e–j (Scheme 2). We performed the polymerization in aqueous d3-acetate buffer at pD~5, conditions where both imines and acylhydrazones are efficiently formed to generate biodynamers. Moreover, we followed the polycondensation by monitoring the signals from the aldehyde group with 1H-NMR spectroscopy, and calculated the consumption of dialdehyde 1 (Table 1). The decreased reactivity of dipeptide hydrazides compared with that of the amino acid hydrazides leads incomplete consumption of the dialdehyde.
Figure 1. Formation of poly(1-e), poly(1-f) and poly(1-g) in aqueous d3-acetate buffer at pD 5. Percentage of unreacted dialdehyde 1 vs time.
Through comparing the consumption of dialdehyde 1 in formation of poly(1-e), poly(1-f) and poly(1-g) (Table 1), we observed that: (1) aromaticity of the side chain plays the most essential role in facilitating polycondensation, which demonstrates the importance of π−π-stacking interactions. As poly(1-h) and poly(1-i) contain an aromatic side chain, more dialdehyde 1 was consumed than for poly(1-f) and poly(1-g); (2) the presence of hydroxyl groups in the side chain has a weaker influence and favors the formation of poly(1-g) via hydrogen bonds, which is also confirmed by comparing poly(1-h) with poly(1-i) in terms of the consumption of
0 100 200 300 400 500 600 700 800 900 10 20 30 40 50 60 70 80 90 100 % Diald e h y d e 1 Time / min poly(1-e) poly(1-f) poly(1-g)
dialdehyde 1; (3) a high density of positive charge (poly(1-f)) has a minor effect and blocks polymerization through electrostatic repulsions between side chains; we observed an increment in dialdehyde 1 consumption for poly(1-h) and poly(1-j) compared to poly(1-f). Meanwhile, we investigated the influence of side chains on the rate of polymerization by monitoring the consumption of dialdehyde 1 in polycondensation (Figure 1, Table 6). Each of the amino acid hydrazides e–g was added to an equimolar amount of dialdehyde 1 to generate the corresponding polymer, and the consumption of dialdehyde 1 was monitored by 1H-NMR spectroscopy until the spectra no longer changed after 2 d. The generation of poly(1-e) is completed in 6 h (Figure 1, Table 6), which suggests that aromatic rings have the most important influence in accelerating the process of polymerization through π−π-stacking interactions. Poly(1-g) reached equilibrium in 1d, which indicates that hydroxyl group does not appear to play an important role in accelerating the rate of polymerization. In the formation of positively charged poly(1-f), signals from dialdehyde 1 were still visible after one week, which illustrates that a high density of positive charge blocks polymerization through electrostatic repulsions. Taken together, these finding are in agreement with the conclusions we drew from monitoring the consumption of dialdehyde 1.
We investigated the morphologies of resulting biodynamers through LS (Table 2 and 5) and cryo-TEM (Table 2 and Figure 2), given that mass spectrometry does not provide information on the length of the intact biodynamers due to the inherent lability of the imine linkages. We observed that poly(1-e) is stabilized by
π−π-stacking interactions and has the biggest particle size and aggregation number,
while poly(1-i) also has the biggest particle size and aggregation number due to the contribution ofπ−π-stacking interactions and hydrogen bonds. Furthermore, poly(1-g) have medium-sized particle and aggregation number through hydrogen-bonding interactions, whereas the sizes of poly(1-f), poly(1-h) and poly(1-j) decreased a lot by virtue of the presence of positive charges. These findings are in line with the conclusions from our analysis of the 1H-NMR spectra.
109 Figure 2. Cryo-TEM images of (a) poly(1−e); (b) poly(1-f); (c) poly(1-g); (d) poly(1-h); (e) poly(1-i); (f) poly(1-j). No stain was used and image acquisition was achieved at a 2 μm defocus. Scale bar = 50 nm.
Table 2. Structural parameters obtained from cryo-TEM and LS. Sample R (nm)a) Rh (nm) b) Rh (nm) c) Mdimer (g/mol) MW,app (g/mol)=KC/I(0) Aggregation number poly(1-e) 65±45 20.04 22.10 787.9 618276 785 poly(1-f) 5.03±1.08 4.04 5.20 770.0 11739 15 poly(1-g) 9.98±1.33 7.94 8.42 687.8 252012 366 poly(1-h) 2.81±0.35 2.29 2.64 828.0 16265 20 poly(1-i) 11.78±2.09 20.42 20.4 737.9 579842 786 poly(1-j) 5.96±1.17 4.63 5.39 728.9 50287 69 poly(2-a) 1.92±0.31 0.68 1.8 329.4 4525 14 poly(2-b) 2.52±0.23 1.22 1.75 320.4 4716 15 poly(2-d) 3.47±0.25 -e) -e) 279.3 5150 18 poly(2-b-c) 1.70±0.18 -e) 1-1.5 649.7f) 6574 10 poly(2-e) 10.19±0.78 1.1 -e) 466.5 27188 58 poly(2-f) 1.51±0.19 0.93 0.67 and 9.35 448.6 22249 50 poly(2-g) 7.13±0.43 -d) -d) 366.4 -d) -d) poly(2-h) 3.10±0.37 3.62 5.18 and 268 506.6 36220 71 poly(2-i) 2.75±0.24 ＜1 -e) 416.5 3244 8 poly(2-j) 2.85±0.29 1.07 0.45 and 13.3 407.5 15295 38 poly(1-2-a) 2.86±0.33 1.29 3.37 980.2f) 18203 19 poly(1-2-d) -d) 26.10 28.6 880.0f) 492996 560 poly(1-2-e) -d) 23.31 23.9 1254.4f) 568220 453 poly(1-2-f) -d) -e) 2 1218.6f) 2933 2-3 poly(1-2-g) -d) 22.05 24.2 1054.2f) 684398 649
Mdimer= dimer molecular weight, Mw = weight-averaged molecular weight, Rg = radius of gyration, Rh = apparent hydrodynamic radius.a) Obtained from cyro-TEM experiments; b) Obtained from SLS measurements and by applying the cumulant method to our data; c) Obtained from SLS measurements and by applying the Contin method to our data; d) Experiments were not performed; e)Signal was very low; f)Mw
of tetramer.
6.2.3 Generation and characterization of biodynamers containing dialdehyde 2 Dialdehyde 2 was introduced into biodynamers to replace dialdehyde 1 and improve the biocompatibility. We prepared the corresponding biodynamers by polycondensation of dialdehyde 2 with monomeric hydrazides a–j (Scheme 3) in mildly acidic aqueous d3-acetate buffer at room temperature and followed by 1H-NMR
111 spectroscopy. Similarly, we calculated the consumption of dialdehyde 2 (Table 3). Unfortunately, we observed that only poly(2-a), poly(2-e) and poly(2-h) formed and consumed most of dialdehyde 2 at room temperature, whereas the other hydrazides only consumed a small amount of dialdehyde 2 at equilibrium. Such findings can be attributed to the decreased reactivity of dialdehyde 2 compared with dialdehyde 1. However, it also demonstrates the importance of aromatic main chains for the formation of biodynamers. Such an aromatic core provides the driving force for polymerization and stabilizes the resulting dynamic proteoids through π−π-stacking interactions. On the other hand, the successful generation of poly(2-a), poly(2-e) and poly(2-h) at room temperature also demonstrates the importance of aromatic side chains in polycondensation through π−π-stacking interactions. To generate the unformed biodynamers, we proposed to enhance the extent of polymerization by increasing reaction temperature or adding dialdehyde 1.
H2N H N N H O O R2 R1 NH2 pD = 5 + N H NH2 H2N O R O OH HO N n + pD = 5 + N H NH2 H2N O R + pD = 5 N N N O 6 H N O R N n O HO N H N N H N O N R O OH HO N n N H N O N H R2 N O R1 N O R
0.5 eq. 0.5 eq. 1 eq. c)
b) a)
OH
2
a, b, d poly(2-a), poly(2-b), poly(2-d)
2
e-j poly(2-e) to poly(2-j)
1 2 a, d poly(1-2-a), poly(1-2-d) pD = 5 N N N O 6 H N O R2 N H n d) poly(1-2-e) to poly(1-2-g) + 0.5 eq. 0.5 eq. 1 2 H2N H N N H O O R2 R1 NH2 e-g 1 eq. + N O R1 O HO OH N H N N H O R2 N O R1 O OH HO N + pD = 5 HN N O N 2 b + c 0.5 eq. 0.5 eq. 1 eq. e) O N N H HO OH N O H2N NaO O n poly(2-b-c)
Scheme 3. Schematic representation of the preparation of dynamic proteoids through reversible polycondensation of (a) dialdehyde 2 with amino acid hydrazides a, b, d; (b) dialdehydes 2 with dipeptide hydrazides e–j; (c) dialdehydes 2 (1.0 eq.) with amino acid hydrazides b (0.5 eq.) and c (0.5 eq.); (d) dialdehydes 1 (0.5 eq.) and 2 (0.5 eq.)
with amino acid hydrazides a or d (1.0 eq.); (e) dialdehydes 1 (0.5 eq.) and 2 (0.5 eq.) with dipeptide hydrazides e–g (1.0 eq.).
Table 3. Consumption of dialdehyde 2 obtained by monitoring the signals from the aldehyde group by 1H-NMR spectroscopy (the integration of the signals from the aldehyde group at 5 min is set to 100%).
Samples Consumption of dialdehyde
2 at room temperature (%) Consumption of dialdehyde 2 at 50 °C (%) poly(2-a) 89 - poly(2-b) 7 - poly(2-d) 4 60 poly(2-e) 85 - poly(2-f) 17 71 poly(2-g) 26 61 poly(2-h) 86 - poly(2-i) 29 81 poly(2-j) 32 62 poly(2-b-c) 8 70
As imine formation can be accelerated by raising the reaction temperature, we performed polycondensations at a higher temperature (50 °C) and observed significant increments in consumption of dialdehyde 2 in all cases. This leads to the formation of the corresponding biodynamers with improved biocompatibility, and demonstrates that higher temperature speeds up polycondensation and facilitates the generation of biodynamers. We designed poly(2-b-c) bearing oppositely charged side chains to evaluate the importance of electrostatic attractions. As poly(2-b-c) cannot form at room temperature, it demonstrates that electrostatic attractions between oppositely charged monomers are not as strong enough to initiate polycondensation as aromatic rings. Compared with poly(2-d) at 50 °C, however, the extent of polymerization of poly(2-b-c) was increased by the electrostatic attractions (Table 3).
We characterized the structures of the dynamic proteoids formed by LS (Table 5 and Figure 5, 6) and cryo-TEM (Figure 3 and 4), and determined their sizes (Table 2). By analogy to biodynamers containing dialdehyde 1, we observed that poly(2-e) has the biggest particle size due to π−π-stacking interactions. Additionally, poly(2-g) bearing hydroxyl groups is stabilized through hydrogen-bonding interactions and has a particle size of 7.13±0.43 nm. On the other hand, poly(1-2-a), poly(1-2-d), poly(1-2-e), poly(1-2-f) and poly(1-2-d) were prepared through polymerization of half an equivalent of both dialdehydes 1 and 2 with one equivalent of the corresponding
113 amino acid hydrazide at room temperature. In case of poly(1-2-d) and poly(1-2-g), 51% of dialdehyde 2 was consumed in comparison with 4% for poly(2-d), and 53% of dialdehyde 2 was consumed compared with 26% for poly(2-g). Whereas no obvious enhancement was observed in case of poly(1-2-a), poly(1-2-e) and poly(1-2-f). It demonstrates that, in some cases, addition of dialdehyde 1 may stabilize the generated biodynamer through π−π-stacking interactions and result in significant enhancement of product yield. From LS and the cryo-TEM image of poly(2-a) (Figure 3), we observed globular nano-object structures with a particle size of 2.86±0.33 nm, which can be ascribed to π−π-stacking interactions between the aromatic cores of dialdehyde 1 and the aromatic side chains of monomer a. In addition, hydroxyl groups in dialdehyde 2 may also contribute to the resulting architectures through hydrogen-bonding and OH−π interactions. In addition, we observed bigger well-ordered nanostructures and aggregation numbers of poly(1-2-d), poly(1-2-e) and poly(1-2-g) than corresponding poly(2-d), poly(2-e) and poly(2-g), which demonstrates that the addition of dialdehyde 2 provides further stability to the resulting biodynamers and facilitates the formation of well-ordered nanostructures through π−π-stacking interactions. Taken together, biodynamers with improved biocompatibility and well-ordered structures can be generated through both methods, which are stabilized by self-folding of the resulting polymers through various non-covalent interactions, including π−π-stacking, hydrogen-bonding and OH−π interactions.
Table 4. Consumption of dialdehydes 1 and 2 obtained by monitoring the signals from the aldehyde group by 1H-NMR spectroscopy (the integration of the signals from the aldehyde group at 5 min is set to 100%).
Samples Consumption of dialdehyde 1 (%) Consumption of dialdehyde 2 (%)
poly(1-2-a) 100 93
poly(1-2-d) 100 51
poly(1-2-e) 100 69
poly(1-2-f) 43 24
Figure 3. Cryo-TEM images of (a) poly(2−a); (b) poly(2−b); (c) poly(2−d); (d) poly(2−b−c); (e) poly(1-2−a). No stain was used and image acquisition was achieved at a 2 μm defocus. Scale bar = 50 nm.
115 Figure 4. Cryo-TEM images of (a) poly(2−e); (b) poly(2-f); (c) poly(2-g); (d) poly(2-h); (e) poly(2-i); (f) poly(2-j). No stain was used and image acquisition was achieved at a 2 μm defocus. Scale bar = 50 nm.
6.3 Conclusions
We designed and prepared a range of dynamic proteoids with well-ordered structures, based on the polycondensation of different types of amino acid and dipeptide hydrazides with nonbiological aromatic dialdehyde 1 and biological aliphatic dialdehyde 2, through formation of two types of reversible C=N bonds (imine and acylhydrazone). By introducing biological dialdehyde 2, the biocompatibility of the resulting biodynamers is enhanced. The use of dialdehyde 2, however, decreases the extent of polymerization, which demonstrates that polycondensation is driven by the self-organization/folding of the resulting polymers mainly through π−π-stacking interactions between the aromatic cores of dialdehyde 1. Thus, we improved the extent of the reaction between hydrazides with dialdehyde 2 by raising the reaction temperature or adding aromatic dialdehyde 1. By using
1
H-NMR spectroscopy and cryo-TEM, we characterized polymerization and the structures of the biodynamers formed and demonstrated that the side chains of amino acid hydrazides, including aromaticity, charge and polarity, have a strong influence on polymerization and structure of the resulting biodynamers. We evaluated the respective importance of the three factors and demonstrated that aromaticity of the side chains plays the most essential role in facilitating polycondensation through
π−π-stacking interactions, hydroxyl groups in the side chains have a less important
influence and also stabilize the architectures formed via hydrogen bonds, whereas a high density of positive charge hinders the generation of biodynamers owing to the electrostatic repulsions. Taken together, these findings provide tools for rational design and synthesis of well-ordered architectures and adaptive dynamic proteoids. The biodynamers generated combine biocompatibility and functionality of biological components with adaptability stemming from dynamic covalent bonds, to achieve synergistic properties. Such dynamic biomaterials hold great potential to be used as functional dynamic biomaterials in both biomedical and bio-engineering fields.
6.4 Experimental section
6.4.1 Synthesis of dialdehydesThe dialdehydes were prepared according to the procedure reported in the literatures (Scheme 4).[4–5]
117 N H Br Br N H CHO OHC iii. 6 O OH 6 O OTs 3 O OTs 3 O OH N O O O 6 iv. v. 1 5 4 8 7 3 6 i. ii. O OH HO OH OH HO HO O OH HO OMe OMe MeO MeO O OBz BzO O OBz BzO OH HO OTr TrO O OH HO OTr TrO O OAc AcO OAc AcO O OH HO NH2 OH HO HCl
vi. vii. ix.
x. O OH HO OH HO viii. xi. xii. 9 10 11 12 2 16 14 13 O OBz BzO OMe OMe MeO MeO xiii. 15 a) b) c) 84% 68% 42% 61% 33% 31% 84% 67% 38% 37% 97% 71% 98%
Scheme 4. Synthesis of monomers 1 and 2. Reagents and conditions: i, NaOH, p-TsCl, H2O, THF;[5a] ii, triethylene glycol, KOH, 100 ˚C;[5a] iii, p-TsCl, TEA, DCM;[5b] iv,
BuLi, DMF, -78 ˚C;[5b] v, 6, Na2CO3, NaI, DMF, 80 ˚C;[5b] vi, (1) NaNO2, AcOH, H2O,
0 ˚C, (2) NaBH4, H2O, dry ice, (3) pyridine, Ac2O;[5c] vii, NaOMe, MeOH;[5c] viii,
TrCl, pyridine;[5d] ix, BzCl, pyridine;[5d] x, AcOH, H2O, 80 ˚C;[5d] xi, (1) TEMPO,
TCCA, DCM, (2) CH(OMe)3, MeOH, p-TsOH; xii, NaOMe, MeOH;[4] xiii, DCl, 50
˚C, D2O.[4]
6.4.2 Synthesis and characterization of dipeptide hydrazides
H N OH O 2 eq HBTU , 4 eq DIPEA DMF, RT, 1h* 2 2 eq Fmoc-L-AA1-OH H N O O H N O Fmoc R1 20% Piperidine/DMF 30 min H N O O NH2 O R1 H N O O H N O R1 O N H R2 1. TFA:H2O/95:5, 1h 2 eq HBTU , 4 eq DIPEA DMF, RT, 1h* 2 2 eq Fmoc-L-AA1-OH Fmoc 2. 20% Piperidine/DMF 30 min H N O O H N O R1 O NH2 R2 5% NH2NH2 in DMF, 2h H2NHN H N O R1 O NH2 R2
Scheme 5. Solid-phase synthesis of dipeptide hydrazides (e–j).
General procedure: Dipeptide hydrazides were prepared by Fmoc standard
solid-phase peptide synthesis (SPPS). In brief, 20% (v/v) piperidine/DMF was used for Fmoc deprotection. Fmoc-protected amino acids were linked progressively to the 4-(Hydroxymethyl)benzoyl-aminomethyl polystyrene (HMBA-AM) resin in a DMF (peptide grade) solution containing HBTU, HOBt, and DIEA, using 10% ninhydrin in
methanol for monitoring the coupling efficacy. Trifluoroacetic acid (TFA)/H2O 95:5
was used for deprotection of side chains. Cleavage from the resin was achieved with 5% hydrazine monohydrate in DMF for 1 h at room temperature. The solvent was evaporated, the residue redissolved in TFA and precipated in cold diethylether. The precipatation was repeated once. Finally, the filtrate was redissolved in H2O and
lyophilized. All the dipeptide hydrazides were purified with preparative RP-HPLC. RP-HPLC conditions used: column, Xterra Prep MS C18, 10 µm, 7.8 x 150 mm, flow rate, 1.0 mL min-1, wavelength 220 nm.
His-His-NH2NH2 (e). Yield: 64%. [α]D20 = +0.018 deg cm3 g−1 (c = 2 mg cm−3, H2O);
mp 80–81 °C; 1H NMR (400 MHz, D2O, δ) 8.74–8.60 (m, 2H), 7.43–7.28 (m, 2H),
4.62 (t, J = 7 Hz, 1H), 4.30 (t, J = 7 Hz, 1H), 3.45–3.32 (m, 2H), 3.28–3.12 (m, 2H);
13
C NMR (101 MHz, D2O, δ) 172.3, 171.2, 137.3, 136.7, 130.5, 128.5, 121.5, 120.4,
54.8, 54.3, 29.2, 21.0; HRMS (ESI) m/z: [M + H]+ calculated for C12H29N8O2,
307.1626; found: 307.1632.
Lys-Lys-NH2NH2 (f). Yield: 41%. [α]D20 = –0.002 deg cm3 g−1 (c = 2 mg cm−3, H2O);
mp 64–66 °C; 1H NMR (400 MHz, D2O, δ) 4.28 (t, J = 7 Hz, 2H), 4.04 (t, J = 7 Hz,
1H), 3.01 (q, J = 8 Hz, 4H ), 1.98–1.78 (m, 4H), 1.76–1.66 (m, 4H), 1.51–1.38 (m, 4H); 13C NMR (101 MHz, D2O, δ) 169.7, 162.7, 52.6, 52.1, 39.0, 38.9, 30.3, 30.1,
26.2, 26.1, 21.9, 21.0; HRMS (ESI) m/z: [M + H]+ calculated for C12H29N6O2,
289.2347; found: 289.2350.
Ser-Ser-NH2NH2 (g). Yield: 47%. [α]D20 = –0.018 deg cm3 g −1
(c = 2 mg cm−3, H2O);
mp 117–118 °C; 1H NMR (400 MHz, D2O, δ) 4.22 (t, J = 4 Hz, 2H), 4.03 (t, J = 6 Hz,
2H), 3.93 (d, J = 4 Hz,1H), 3.86 (d, J = 6 Hz, 1H); 13C NMR (101 MHz, D2O, δ)
168.7, 163.8, 63.2, 61.2, 60.5, 57.3; HRMS (ESI) m/z: [M + H]+ calculated for C6H15N4O4,207.1088; found: 207.1090.
Trp-Lys-NH2NH2 (h). Yield: 67%. [α]D20 = +0.038 deg cm3 g−1 (c = 2 mg cm−3,
H2O); mp 107–108 °C; 1H NMR (400 MHz, D2O, δ) 7.56 (dd, J = 16, 8 Hz, 2H), 7.32
(s, 1H), 7.28 (t, J = 7 Hz, 1H), 7.17 (t, J = 8 Hz, 1H), 4.26 (t, J = 8 Hz, 1H), 4.14 (t, J = 7 Hz, 1H), 3.39 (d, J = 7 Hz, 2H), 2.93 (t, J = 8 Hz, 2H), 2.93 (t, J = 8 Hz, 2H), 1.73–1.54 (m, 4H); 13C NMR (101 MHz, D2O, δ) 170.6, 169.1, 136.1, 126.5, 125.1,
121.9, 119.4, 117.9, 111.8, 106.1, 53.6, 51.7, 39.0, 30.7, 26.6, 26.2, 21.6; HRMS (ESI) m/z: [M + H]+ calculated for C17H27N6O2, 347.2190; found: 347.2195.
119 His-Ser-NH2NH2 (i). Yield: 57%. [α]D20 = –0.008 deg cm3 g
−1
(c = 2 mg cm−3, H2O);
mp <50 °C; 1H NMR (400 MHz, D2O, δ) 8.72 (s, 1H), 7.45 (s, 1H), 4.49 (t, J = 7 Hz,
1H), 4.37 (t, J = 7 Hz, 1H), 3.89–3.80 (m, 2H), 3.48–3.41 (m, 2H); 13C NMR (101 MHz, D2O, δ) 168.2, 162.2, 133.3, 127.3, 117.8, 61.4, 59.9, 56.8, 28.8; HRMS (ESI)
m/z: [M + H]+ calculated for C9H7N6O3, 257.1357; found: 257.1359.
Lys-Ser-NH2NH2 (j). Yield: 45%. [α]D20 = –0.016 deg cm3 g−1 (c = 2 mg cm−3, H2O);
mp 56–58 °C; 1H NMR (400 MHz, D2O, δ) 4.34 (t, J = 7 Hz, 1H), 4.17 (t, J = 5 Hz,
1H), 3.93–4.07 (m, 2H), 3.00 (t, J = 7 Hz, 1H), 1.97–1.78 (m, 2H), 1.76–1.64 (m, 2H), 1.55–1.39 (m, 2H); 13C NMR (101 MHz, D2O, δ) 169.7, 163.0, 60.8, 54.0, 52.7, 38.9,
30.2, 26.2, 20.9; HRMS (ESI) m/z: [M + H]+ calculated for C9H22N5O3, 248.1717;
found: 248.1720.
6.4.3 Preparation of dynamic proteoids
Preparation of poly(1-e), poly(1-f), poly(1-g), poly(1-h), poly(1-i) and poly(1-j) Monomer 1 (20 mM in D2O, 0.25 mL), dipeptide hydazide (20 mM in D2O, 0.25
mL), aqueous d3-acetate buffer (400 mM, 0.25 mL, pD 5) and D2O (0.25 mL) were
rapidly mixed, and the pD was set to 5 by addition of DCl (1.0 M) or NaOD (1.0 M) and left to stand at 25 °C. For LS measurements, samples were passed through a 200 nm syringe filter immediately after mixing.
Preparation of poly(2-a) to poly(2-j)
Monomer 2 (20 mM in D2O, 1 mL) was added aμ DCl solution (35% in D2O by
weight). The mixture was stirred at 50 °C for 24 h and neutralized with NaOD (40% in D2O by weight) to give solution A. Solution A (0.25 mL), dipeptide hydrazide (20
mM in D2O, 0.25 mL), aqueous d3-acetate buffer (400 mM, 0.25 mL, pD 5) and D2O
(0.25 mL) were rapidly mixed, and the pD was set to 5 by addition of DCl (1.0 M) or NaOD (1.0 M) and left to stand at 25 °C or 50 °C. For LS measurements, samples were passed through a 200 nm syringe filter immediately after mixing.
Preparation of poly(2-b-c)
Solution A (0.25 mL), monomer b (20 mM in D2O, 0.25 mL), monomer c (20 mM
in D2O, 0.25 mL) and aqueous d3-acetate buffer (400 mM, 0.25 mL, pD 5) were rapidly mixed, and the pD was set to 5 by addition of DCl (1.0 M) or NaOD (1.0 M) and left to stand at 50 °C. For LS measurements, samples were passed through a 200 nm syringe filter immediately after mixing.
Preparation of poly(1-2-a) and poly(1-2-d)
Monomer 1 (20 mM in D2O, 0.25 mL), solution A (0.25 mL), dipeptide hydrazide
(20 mM in D2O, 0.25 mL) and aqueous d3-acetate buffer (400 mM, 0.25 mL, pD 5) were rapidly mixed, and the pD was set to 5 by addition of DCl (1.0 M) or NaOD (1.0 M) and left to stand at 25 °C. For LS measurements, samples were passed through a 200 nm syringe filter immediately after mixing.
6.4.4 Light scattering methods
The measurements used a 3D dynamic light scattering (DLS) spectrometer (LS Instruments, Fribourg, Switzerland) equipped with a 25 mW HeNe laser (JDS uniphase) operating at =632.8 nm, a two channel multiple tau correlator (1088 channels in autocorrelation), a variable-angle detection system, and a temperature-controlled index matching vat (LS Instruments). The scattering spectrum was measured using two single mode fibre detections and two high sensitivity APD detectors (Perkin Elmer, model SPCM-AQR-13-FC).
Fluctuations in the scattered intensity with time I(q,t) (also called count rate), measured at a given scattering angle or equivalently at a given scattering wave vector q=(4n/)sin(/2), are directly reflecting the so-called Brownian motion of the scattering particles. In dynamic light scattering (DLS), the fluctuation pattern is translated into the normalized time autocorrelation function of the scattered intensity,
g(2)(q,t) defined as: 2 ) 2 ( ) 0 , q ( I ) t , q ( I ) 0 , q ( I ) t , q ( g (1)
It is related to the so-called dynamic structure factor (or concentration fluctuations autocorrelation function), g(1)(q,t), via the Siegert relation:
𝑔(2)(𝑞, 𝑡) − 1 = 𝛽|𝑔(1)(𝑞, 𝑡)|2 (2)
Where is the coherence factor, which in our experiments is varying between 0.5 and 0.8, depending on the samples and the setup geometry. The normalized dynamical correlation function, g(1)(q,t), of concentration fluctuations is defined as:
2 ) 1 ( ) 0 , q ( c ) t , q ( c ) 0 , q ( c ) t , q ( g (3)
Where c(q,t) and c(q,0) represent fluctuations of the concentration at time t and
zero, respectively. The distribution of decay rates G(Г) was determined using the CONTIN algorithm based on the inverse Laplace transform of g(1)(q,t):
121
0 ) 1 ( d ) t exp( ) ( G ) t , q ( g (4)For a diffusive process, with characteristic time, τ=1/Г, inversely proportioned to q²,
g(1)(q,t)~exp(-Dq2t), with D the mutual diffusion coefficient. The Stokes-Einstein
relation allows one to determine the hydrodynamic radius RH of the scattered objects;
RH=kT/6πηD, if the temperature T and solvent viscosity are known (here =1.002
cP at 20 °C for water).
In static light scattering (SLS) experiments, the excess of scattered intensity is measured with respect to the solvent. The so-called excess Rayleigh ratio was deduced using a toluene sample reference for which the excess Rayleigh ratio is well-known (Rtoluene=1.352210-5 cm-1 at 633 nm): 𝑅𝑠𝑜𝑙𝑢𝑡𝑒(𝑐𝑚−1) = 𝐼𝑠𝑜𝑙𝑢𝑡𝑖𝑜𝑛−𝐼𝑠𝑜𝑙𝑣𝑒𝑛𝑡 𝐼𝑡𝑜𝑙𝑢𝑒𝑛𝑒 × ( 𝑛 𝑛𝑡𝑜𝑙𝑢𝑒𝑛𝑒) 2 × 𝑅𝑡𝑜𝑙𝑢𝑒𝑛𝑒 (5)
The usual equation for absolute light scattering combines the form factor P(q), the structure factor S(q) and the weight-average molecular weight Mw of the scattered
objects: ) ( ) ( ) ( ² 4 ) ( 2 4 0 2 q S q P cM dc dn N n q R w a (6)
where K=42n2(dn/dC)2/NA4 is the scattering constant (refractive index n=1.33 at 20 °C) and NA the Avogadro’s number. A refractive index increment of dn/dC = 0.185 cm3/g was considered to be a sufficient approximation.3 A classical Guinier analysis was used to determine Mw as well as the radius of gyration, Rg, for particles larger
than 20 nm: 1 𝑅(𝑞)⁄ = 1 𝑅(0)⁄ (1 +𝑞
2𝑅
𝑔 2
Figure 5. Intensity correlation function obtained from dynaimic light scattering (DLS) experiments. The inset represents the size distribution obtained by applying the Contin method to the data.
123 Table 5. Structural parameters obtained from LS.
Sample Rh (nm)a) Polydispers ity, k2/k12 a) Rh (nm) b) Rg (nm) I(q2=0) (a.u.) = I/Itol(q=0) (cm-1) Mdimer (g/mol) MW,app (g/mol)= KC/I(0) Aggregatio n number Cdimer or trimer (g/cm310-3) poly(1-e) 20.04 0.13 22.1 0 48 56 787.9 618276 785 3.9395 poly(1-f) 4.04 0.20 5.20 -d) 1.152 770.0 11739 15 3.85 poly(1-g) 7.94 0.10 8.42 -d) 20 687.8 252012 366 7.94 poly(1-h) 2.29 0.15 2.64 -d) 1.66 828.0 16265 20 4.14 poly(1-i) 20.42 0.13 20.4 54.3 49.2 737.9 579842 786 3.6895 poly(1-j) 4.63 0.094 5.39 -d) 4.32 728.9 50287 69 3.6445 poly(2-a) 0.68 0.33 1.8 -d) 0.286 329.4 4525 14 1.647 poly(2-b) 1.22 0.43 1.75 -d) 0.29 320.4 4716 15 1.602 poly(2-d) -d) -d) -d) -d) 0.28 279.3 5150 18 1.3965 poly(2-b-c) -d) -d) 1-1.5 -d) 0.36 649.7e) 6574 10 1.62425 poly(2-e) 1.1 -d) -d) -d) 1.57 466.5 27188 58 2.3325 poly(2-f) 0.93 -d) 0.67, 9.35 -d) 1.26 448.6 22249 50 2.243 poly(2-g) -c) -c) -c) -c) - c) -c) 366.4 -c) -c) poly(2-h) 3.62 0.66 5.18, 268 -d) 2.22 506.6 36220 71 2.533 poly(2-i) ＜1 -d) -d) -d) 0.27 416.5 3244 8 2.0825 poly(2-j) 1.07 -d) 0.45, 13.3 -d) 0.83 407.5 15295 38 2.0375 poly(1-2-a) 1.29 0.33 3.37 -d) 1.14 980.2e) 18203 19 2.4505 poly(1-2-d) 26.10 0.13 28.6 64 25 880.0e) 492996 560 2.2 poly(1-2-e) 23.31 0.146 23.9 54 41 1254.4e) 568220 453 3.136 poly(1-2-f) -c) -d) 2 -d) 0.32 1218.6e) 2933 2-3 3.0465 poly(1-2-g) 22.05 0.127 24.2 62.9 41.5 1054.2e) 684398 649 2.6355
Mdimer= dimer molecular weight, Mw = weight-averaged molecular weight, Rg = radius of gyration, Rh = apparent hydrodynamic radius. a) Obtained from SLS measurements and by applying the cumulant method to our data; b) Obtained from SLS measurements and by applying the Contin method to our data; c) Experiments were not performed; d)Signal was very low; e)Mw of tetramer.
Table 6. Percentage of unreacted dialdehyde 1 vs time obtained by monitoring the signals from the aldehyde group by 1H-NMR spectroscopy.
time(min) poly(1-e) (%) poly(1-f) (%) poly(1-g) (%) 0 100 100 100 15 88.947 94.333 84.309 30 82.502 89.513 71.257 45 75.911 86.388 64.587 60 69.379 84.269 58.685 75 62.351 82.627 54.079 90 54.972 80.35 49.424 105 49.373 77.225 46.113 120 44.882 74.417 41.603 135 38.612 72.405 38.964 150 34.733 71.981 35.988 165 31.175 70.922 33.253 180 27.763 69.915 30.71 195 24.759 68.538 28.263 210 21.931 67.161 26.727 225 19.189 66.261 24.04 240 16.594 65.678 23.033 255 14.377 65.148 21.737 270 11.928 64.619 20.441 285 10.12 62.659 19.434 300 8.661 62.129 18.234 315 7.203 61.758 17.131 330 5.803 61.017 16.315 345 4.579 60.275 14.971 360 3.995 59.852 14.107 375 4.024 59.746 13.388 390 4.083 58.739 12.812 405 3.932 58.104 12.236 420 3.82 57.362 11.708 435 3.995 56.886 11.276 450 3.937 56.515 10.797 465 4.054 56.568 10.365 480 3.762 56.462 9.981 495 3.85 56.303 9.837 510 3.985 56.356 8.877 525 3.997 55.72 8.253 540 3.937 56.144 7.869
125 555 3.991 55.614 7.63 570 3.854 54.237 7.294 585 3.975 54.131 7.006 600 3.937 54.343 6.814 615 3.989 53.602 6.622 630 3.999 53.549 6.286 645 3.827 52.966 6.142 660 3.937 53.443 5.998 675 3.975 51.165 5.758 690 3.82 51.112 5.566 705 3.937 51.218 5.47 720 3.995 51.377 5.518 735 3.851 51.43 5.422 750 3.831 51.112 5.374 765 3.852 50.742 5.471 780 3.859 51.695 5.326 795 3.856 51.377 4.942 810 3.995 50.583 4.972 825 3.937 50.742 4.846 840 3.995 50.477 5.134 855 3.991 50 4.993 870 3.995 51.642 4.99 885 3.829 51.589 4.944 900 3.989 51.271 4.894
6.5 Contributions from co-authors
Cyro-TEM measurements were performed by Dr. M. C. A. Stuart. LS measurements and analysis was performed by Prof. Dr. E. Buhler.
6.6 References
[1] A. K. H. Hirsch, E. Buhler, J.-M. Lehn, J. Am. Chem. Soc. 2012, 134, 4177.
[2] a) Y. Liu, M. C. A. Stuart, E. Buhler, J.-M. Lehn, A. K. H. Hirsch, Adv. Funct. Mater. 2016,
26, 6297; b) D. Zhao, J. S. Moore, J. Am. Chem. Soc. 2003, 125, 16294.
[3] a) C. Godoy-Alcántar, A. K. Yatsimirsky, J.-M. Lehn, J. Phys. Org. Chem. 2005, 18, 979; b) A. Aissaoui, B. Martin, E. Kan, N. Oudrhiri, M. Hauchecorne, J.-P. Vigneron, J.-M. Lehn, P. Lehn, J. Med. Chem. 2004, 47, 5210.
[5] a) M. T. Stone, J. S. Moore, Org. Lett. 2004, 6, 469; b) J. F. Folmer-Andersen, E. Buhler, S.-J. Candau, S. Joulie, M. Schmutz, J.-M. Lehn, Polym. Int. 2010, 59, 1477; c) M. Ohashi, K. Gamo, Y. Tanaka, M. Waki, Y. Beniyama, K. Matsuno, J. Wada, M. Tenta, J. Eguchi, M. Makishima, N. Matsuura, T. Oyama, H. Miyachi, Eur. J. Med. Chem. 2015, 90, 53; d) J. Kuszmann, G. Medgyes, S. Boros, Carbohydr. Res. 2005, 340, 1739.
