Polymer–protein conjugates: an enzymatic activity perspective

Abstract

Proteins have been modified with polymers in diverse manners over the past 30 years. However, while proteins have been used to prepare many functional constructs, they are sensitive biomolecules and their bioactivity can be either positively or negatively influenced by many different aspects of polymer modification. The primary focus of this review article is to highlight the opportunities offered by new trends in protein modification, and specifically how they influence the overall biological activity of the conjugate, including its dependence on temperature and pH. We survey the effect of polymer molecular weight, number of conjugated polymer chains, polymer coupling strategy (including random versus site-specific coupling, “grafting from”, and multi-point covalent attachment), polymer architecture (including branched and comb-type), polymer interactions with the protein (including electrostatic and host–guest interactions), polymer interactions with enzyme substrate, and polymer biodegradability. We have selected six enzymes, which have been extensively modified with polymers in diverse fashions in the literature, as basis for this discussion. These proteins are L-asparaginase, alpha-chymotrypsin, trypsin, lysozyme, bovine serum albumin, and papain. This review includes polymers such as poly(ethylene glycol) (PEG), polysaccharides, polypeptides, and other synthetic (vinyl) polymers. From the discussed literature we attempt to extract tentative general trends observed between state-of-the-art methods of preparing protein–polymer conjugates and the activity of the conjugate.

---

Marc A. Gauthier

Marc A. Gauthier was awarded his Ph.D. in 2007 for his work with T.H. Ellis and X.X. Zhu at the University of Montreal (Montreal, Canada). During this period he received fellowships from the Natural Science and Engineering Council of Canada and the Fonds Québecois de la Recherche sur la Nature et les Technologies (FQRNT). He then pursued post-doctoral research (FQRNT) with H.-A. Klok (EPFL, Switzerland) in the field of peptide/protein–polymer conjugation. He currently holds the position of Research Fellow at the Institute of Pharmaceutical Sciences (IPW) at the Swiss Federal Institute of Technology Zürich (ETHZ) (Zürich, Switzerland), where he is pursuing independent research involving therapeutic protein conjugates.

Harm-Anton Klok

Harm-Anton Klok is Full Professor at the Institutes of Materials and Chemical Sciences and Engineering at the Ecole Polytechnique Fédérale de Lausanne (EPFL) (Lausanne, Switzerland). His current research interests include peptide/protein-based materials and peptide/protein-polymer hybrids, surface-initiated polymerization and polymer brushes, controlled/“living” polymerization and macromolecular engineering as well as dendritic and hyperbranched polymers. Harm-Anton Klok is recipient of the 2007 Arthur K. Doolittle Award of the American Chemical Society and is Associate Editor of the American Chemical Society journal Biomacromolecules.

---

1. Introduction

Background

Over the past 30 years proteins have been extensively modified with either well- or ill-defined polymers by means of either well- or ill-defined coupling chemistry. By far, the desire to overcome the intrinsic limitations of protein therapeutic agents has been the driving force for the developments in this field, given that approximately 25% of all new medicines approved by the American Food and Drug Administration are protein-based.¹ Initially, the philosophy for overcoming the immunogenicity of therapeutic proteins, their renal elimination, and their proteolysis while maintaining therapeutic activity has been to modulate the number, the length, and to a certain extent the architecture (i.e., linear versus branched) of the polymer segments used for preparing the conjugate. Early on, many natural polymers as well as synthetic polymers such as poly(ethylene glycol) (PEG) have been evaluated for this purpose. Possibly due to the fact that PEG was one of the first non-immunogenic, well-defined (hetero)telechelic polymers capable of conveying “stealth” properties to proteins, it is not surprising that the PEGylation (i.e., the covalent attachment of PEG chains) of therapeutic proteins has dominated this field for many years^2,3 and that several therapeutic PEGylated proteins have reached the market.^4–6 More recently, the philosophy for overcoming the shortcomings of therapeutic proteins with minimal detrimental effect on protein biological activity has changed to controlling the site at which the polymer is attached to the protein.^7–9 This has become a possibility with the development of a veritable (bio)chemical toolbox of orthogonal coupling strategies for residue- or site-specific protein modification.^10–22 Concurrently, access to well-defined (hetero)telechelic polymers other than PEG has become possible due to advances in the field of controlled/“living” polymerization.^23–27 These combined developments have lead to a boom in the number of studies implicating protein–polymer conjugates, whether for therapeutic purposes or not. With these advances, many new possibilities are available for adjusting the properties of the final conjugate. One aspect which is of prime importance in the field is that while proteins are used to prepare many functional constructs, they are sensitive biomolecules and their bioactivity can be either positively or negatively influenced by many different aspects of polymer modification. The influence of polymer modification of proteins has previously been discussed by others in the context of PEGylation.^3–6,28,29 However, a discussion which puts these findings into a broader context and which includes modern design and synthetic strategies, inaccessible to classic PEGylation, is lacking. The primary focus of this review article is to highlight the trends observed for these new developments in protein modification, and specifically how they influence the overall biological activity of the conjugate, including its dependence on temperature and pH. This contribution is divided into 6 sections which successively discuss the effect of the number and molecular weight of coupled polymer chains (i.e., lessons learned from PEGylation), polymer coupling strategy, polymer architecture, polymer interaction with the protein, polymer interaction with enzymatic substrate (vide infra), and polymer biodegradability.

Scope

In order to have a reliable, comparable parameter with which to compare the biological properties of protein–polymer conjugates prepared over the past three decades, we have selected enzymes with well-defined and/or commercially available enzymatic substrates for discussion. We have avoided discussing proteins whose activity assays involve live cells, as these tests have more potential to be influenced by unrelated cell- or user-specific variables, which may potentially complicate or bias the establishment of trends within the literature. This implies that very recent and exciting research on conjugates prepared from interleukins, leptin, interferons, antibody fragments, streptavidin, or enzymes with poorly defined enzymatic assays fall outside the scope of this review. Polymer-modification of these types of proteins has been reviewed elsewhere.^7,30,31 Given the vastness of the literature as well as the overwhelming number of publications in this field published every year, we have restricted our discussion to six enzymes. By no means is this contribution intended to be comprehensive and is rather geared towards providing some perspective in the properties of protein–polymer conjugates. The choice of these proteins was subjective but was made based on the criteria that they have been modified in numerous and diverse fashions by synthetic and/or natural polymers. This implies that not all selected proteins are necessarily of therapeutic interest and may simply be popular model proteins for proof-of-concept studies (e.g., owing to cost, availability, or structural considerations) or proteins useful for industrial applications, etc. We have also limited our discussion to water-soluble protein–polymer conjugates, thus eliminating the plethora of studies on enzymes immobilized on insoluble polymer matrices.

The structural and enzymatic parameters of all discussed conjugates are listed in Tables 1–6 and serve as basis for the discussions herein. Enzymatic activity is always given as a percentage of that of the native enzyme and the numerical value is taken directly from the corresponding study, unless preceded by a “∼” which indicates that it was estimated from a figure. When only enzyme kinetic parameters such as k_cat (turn over number) and K_M (Michaelis constant) are given in the original manuscript, these values are used for discussion. From these parameters we make no attempt to calculate an enzymatic efficiency (k_cat/K_M) of the conjugates, as this parameter is ill-suited for comparing enzymes.³²

2. Polymer molecular weight and number of chains

Over the past 30 years, proteins have been extensively modified with PEG in order to establish structure–activity relationships between molecular weight, coupling chemistry, and extent of modification on the activity of the final conjugate. PEGylation is an excellent tool for evaluating the effects of polymer molecular weight and degree of polymer coupling (to proteins) because it does not bear functional side-chains which could potentially interact with the protein and thus bias the results. Classically, the number and length of conjugated polymer chains is thought to correlate with the amount of polymer present on the surface of the enzyme, restricted diffusion of substrate towards the enzymes catalytic site, and reduced bioactivity. In this section we will treat the polymers as being chemically inert and verify the effect of these two parameters on enzyme activity. We will begin by discussing the lessons learned from the extensive history of PEGylation (as demonstrated by the multiple entries in Tables 1–6 involving PEG) and then we will discuss other polymers. This will be followed by a discussion of general trends which can be extracted from the analyzed literature, and which are summarized in Table 7.

Table 1 Structural and enzymatic properties of L-asparaginase-polymer conjugates.^a

Polymer	Residue (number)^b	MW (chain, conjugate)^c	Coupling method	Activity (%)	KM/mM (native)	kcat/s^−1 (native)	Comment, ref.
a ASNase is from E. coli unless otherwise mentioned. b Number of residues modified on protein. c Molecular weight of conjugate only given for multi-site attachment. Unless identified by superscripts d (Mn) or e (Mv), the nature of the average molecular weight was not specified. f L-Asparagine. g Aspartic acid-β-p-nitroanilide. h Not given. i E.carotovora.
Synthetic polymers
MPEG	Lys (18–73)	5 kDa	TCT	6–40^f, 15–50^g	—	—	Effect of molecular weight, ^33
MPEG	Lys (77)	750 Da	TCT	12^f, 20^g	—	—
MPEG	Lys (70)	1.9 kDa	TCT	14^f, 25^g	—	—
MPEG	Lys (50)	5 kDa	NHS	30^f	—	—	Ref. 36
MPEG	Cys (4)	5 kDa	bisAlk	100	—	—	Site-specific^58
MPEG	Cys (4)	10 kDa	bisAlk	100	—	—
MPEG	Cys (4)	20 kDa	bisAlk	100^f	—	—
MPEG2	Lys (33–52)	2 × 5 kDa	TCT	11–30^f	—	—	Branched polymer^34
MPEG2	Lys(52)	2 × 5 kDa	TCT	8^f	∼KM,nat	—	Branched polymer^91
MPEG2	Lys (2.5–30)	2 × 5 kDa	TCT	0–74^h	—	—	Branched polymer^35
MPEG	Lys (35–49)^i	5 kDa^d	NHS	110	3.33 × 10^−3	11.83	Effect of polymer architecture^37,38
MPEG2	Lys (27–37)^i	2 × 5 kDa^d	NHS	130–133^f	3.30 × 10^−3	13
					(3.31 × 10^−3)^f	(8.7)^f
MPEG	Lys (73)	5 kDa	TCT	0.9	—	—	Multi-site attachment. Comb-shaped^74
MPEG2	Lys (52)	2 × 5 kDa	TCT	11	—	—
PM13	Lys (46)	13 kDa, n.c.	MA	45.5	—	—
PM100	Lys (31)	100 kDa, n.c.	MA	85.3^f	—	—
PMPEG5000-g-VP-co-MA)	Lys (30)	77 kDa, n.c.^d	MA	58.6^f	—	—	Multi-site attachment. Comb-shaped^89
P(VP-co-MA)	Lys (63)	34 kDa, n.c.^e	MA	8.3^f	—	—	Multi-site attachment^90
Polypeptides
P(DL-alanine)	Lys (23.5)	5.4 kDa^d	NCA	20	0.011	—	Grafting from^48
P(DL-alanine)	Lys (17)	2.2 kDa^d	NCA	48	—	—
P(DL-alanine)	Lys (31.5)^i	3.5 kDa^d	NCA	65^f	0.03	—
					(0.011)	—
					(0.04)^i	—
Rat albumin	Lys (n.g.)	69 kDa	GA	60^f	2-3 × KM,nat		∼12 chains/enzyme^87
Silk fibroin	Lys (28–55)	40–120 kDa, n.c.	GA	∼40–80^f	0.84 (4.77)	—	Natural polymer^72
Sericin	Lys (45)	20–60 kDa, n.c.	GA	∼25^f	0.159 (10.4)^f	—	Activity for pH = 7.4^93
Carbohydrate polymers
N,O-CM-Chit	Lys (∼10)^b	40, 522 kDa	GA	∼80^f	—	—	Multi-site attachment. ∼10 chains/enzyme^94
Levan	Lys (n.g.)^i	75 kDa	RA	90.5	0.036	—	Multi-site attachment. Hyperbranched polymer. 50, 24, and 40 wt% enzyme in conjugate^73
Levan	Lys (n.g.)^i	75 kDa	RA	76	0.052	—
Levan	Lys (n.g.)^i	2000 kDa, n.c.	RA	58^f	0.095	—
					(0.025)^f	—

---

Table 2 Structural and enzymatic properties of α-chymotrypsin-polymer conjugates

Polymer	Residue (number)^a	MW (chain, conjugate)^b	Coupling method	Activity (%)	KM/mM (native)	kcat/s^−1 (native)	Comment, ref.
a Number of residues modified on protein. b Molecular weight of conjugate only given for multi-site attachment. Unless identified by superscript c (Mn) the nature of the average molecular weight was not specified. d N α-Acetyl-L-phenylalanine ethyl ester. e N α-Benzoyl-L-tyrosine-p-nitroanilide. f GlyGlyPhe-p-nitroanilide. g N α-Benzoyl-L-tyrosine ethyl ester. h Azocasein. i N-trans-cinnamoyl imidazole. j Z-GlyLeuPhe-p-nitroanilide. k GlyValPhe-p-nitroanilide. l Succ-AlaAlaProPhe-p-nitroanilide. m N α-Acetyl-L-tyrosine ethyl ester. n Bovine serum albumin. o Casein.
Synthetic polymers
PHPMA	Lys (n.g.)	90 kDa	TCT	0.5	2	81	Multi-site attachment.
	Lys (n.g.)	53 kDa	Hyd	1.1	2	—	n.c., 3.5, and 24 wt (%)
	Lys (n.g.)	13 kDa	pNP	23.3^d	2	79	Enzyme in conjugate^77
					(1.9)^d	(68.6)^d
MPM-06	Lys (n.g.)	n.g.	CDI	79^e	—	—	Anionic polymer^97
MPEG	Lys (10)	5 kDa	SC	—	0.0298	0.180	Effect of number of chains^42
	Lys (14)	5 kDa	SC	—	0.0442	0.154
					(0.0433)^f	(0.145)^f
PHPMA	Lys (3)	5.5 kDa^c	NHS	—	0.055	0.24	Polymeric substrate also tested^50
	Lys (5)	3.7 kDa^c	NHS	—	0.065	0.27
	Lys (5)	2.7 kDa^c	NHS	—	0.05	0.21
	Lys(3)	2.7 kDa^c	NHS	—	0.045	0.18
	Asp/Glu (7)	2.5 kDa^c	NHS	—	0.3	0.34
					(0.39)^j	(0.83)^j
PHPMA	Lys (13)	2.7 kDa^c	NHS	72–74	1.807	0.348	H-bonding polymer^51
	Lys (13)	5.1 kDa^c	NHS	88	1.612	0.372
	Lys (9)	10.9 kDa^c	NHS	109–112^f^k	1.364	0.424
					(1.052)^f	(0.353)^f
PNIPAAm-co-AADG	Lys (10)	>300 kDa	RA	50	0.17		Multi-site attachment. 14.7, 18.2, and 29.4 wt% enzyme.
	Lys (12)	>300 kDa	RA	28	0.13
	Lys (12)	>300 kDa	RA	31^l	0.13
					(0.12)^l		Variable %AADG in polymers^75
PPEGMA	Lys (1)	n.c., 45.6 kDa^c	ABr	86	—	—	Grafting from. Comb-shaped polymers^53
	Lys (4)	n.c., 52.3 kDa^c	ABr	54	—	—
	Lys (8)	n.c., 90.5 kDa^c	ABr	42	—	—
MPEG	Lys (1)	5, 32.4 kDa	NHS	69	—	—
PPEGMA	Lys (1)	160, 209 kDa^c	NHS	57	—	—
PNaPS	Lys (4)	n.c., 78.1 kDa^c	ABr	46	—	—
PPEGMA475	Lys (4)	n.c., 52.2 kDa^c	ABr	61	—	—
PDMAEMA	Lys (4)	n.c., 44.3 kDa^c	ABr	38^l	—	—
MPEG	Lys (0.7–8)	5 kDa	NHS	50–85^l	0.025–0.084		Effect of number of chains^41
					(0.013)^l
MPEG	Lys (4–8)	700 Da	NHS	—	0.07–0.015	6.2–11.6	Effect of molecular weight^40
	Lys (1–9)	2 kDa	NHS	—	0.08–0.19	7.6–13.5
	Lys (1–8)	5 kDa	NHS	–	0.08–0.11	8.3–13.1
					(0.05)^l	(8.6)^l
Carbohydrate polymers
PSucr	Lys (8–11)	400 kDa, n.c.	RA	55–64^m, 35^n	—	—	Multi-site attachment. Hyperbranched polymers^78
PSucr	Lys (8–11)	70 kDa, n.c.	RA	65–75^m, 26^n	—	—
CMC	Lys (11–13)	12 kDa, n.c.	RA	77–80^m,80^n	—	—
Dextran	Lys (13,14)	250 kDa, n.c.	RA	50–53^m, 29^n	—	—
CM-PβCD	Lys (4)	13 kDa	CDI	115^m,105^o	—	—	26 wt% enzyme^98
Dextran	Lys (1–8)	10 kDa	NHS	—	0.044–0.052	4.3–9.4	Single-point^55
					(0.046)^l	(9.9)^l

---

Table 3 Structural and enzymatic properties of trypsin-polymer conjugates.^a

Polymer	Residue (number)^b	MW(chain, conjugate)^c	Coupling method	Activity (%)	KM/mM (native)	kcat/s^−1 (native)	Comment, ref.
a Trypsin from Bovine pancreas unless otherwise mentioned. b Number of residues modified on protein. c Molecular weight of conjugate only given for multi-site attachment. Unless identified by superscripts d (Mn) or e (Mw) the nature of the average molecular weight was not specified. f N α-Tosyl-arginine methyl ester. g Casein. h N α-Benzoyl-DL-arginine-p-nitroanilide. i N α-Benzoyl-L-arginine-ethyl ester.
Synthetic polymers
Activated PVP	Lys (n.g.)	10, 150 kDa	NHS	173^h, 20^g	—	—	Multi-site attachment^103
MPEG	Lys (3)	5 kDa^d	TCT	95	—	—	Effect of number of chains^45
	Lys (8,9)	5 kDa^d	TCT	150^i	—	—
MPM-06	Lys (n.g.)	n.g.	CDI	70^h	—	—	Anionic polymer^97
MPEG	Lys (12–14)	350 Da	NPC	154^i, 366^h	0.68	10.0	Effect of molecular weight^43
	Lys (12–14)	550 Da	NPC	145^i, 417^h	0.62	8.7
	Lys (12–14)	750 Da	NPC	145^i, 390^h	0.46	8.6
	Lys (12–14)	2 kDa	NPC	154^i, 355^h	0.41	7.2
	Lys (12–14)	5 kDa	NPC	154^i, 364^h	0.42	6.5
	Lys (12–14)	8 kDa	NPC	127^i, 293^h	0.49	8.2
MPEG2	Lys (12–14)	2 × 5kDa	TCT	127^i, 166^h	0.37	2.4
					(0.80)^h	(2.5)^h
PCVBPC	Lys (6,7)	18 kDa	AzoT	17	—	—	Grafting from^68
P(MAA-co-MMA)	Lys (6,7)	8.5 kDa	AzoT	14^g	—	—
MPEG2	Lys (7,8)	2 × 5 kDa^d	NHS	120	0.076	29.8	Branched polymer^37
	Lys (8,9)	2 × 5 kDa^d	NHS	125^f	0.080	38.5
					(0.082)^f	(13.8)^f
PNIPAAm-co-GEMA	Lys (12.9)	37.1 kDa	CDI	23.3	—	—	Thermosensitive polymer^104
	Lys (13.8)	40.5 kDa	CDI	27.2	—	—
	Lys (14.2)	37.8 kDa	CDI	14.9^h	—	—
PNIPAAm	Lys (1–12)	5 kDa^d	NHS	95–124^f	<KM,nat^h	>kcat,nat^h	Effect of number of chains^52
PDMAPS	Lys (2.4)	11 kDa	CDI	∼100^h	<KM,nat	—	Ionic polymer^95
MPEG	Lys (4,5)	1.1 kDa	CDI	96	1.15	6.6	Effect of coupling chemistry. Effect of molecular weight^46
	Lys (4,5)	2 kDa	CDI	77	0.93	5.3
	Lys (5,6)	5 kDa	CDI	65	0.79	4.0
	Lys (8,9)	1.1 kDa	TCT	18	0.83	1.1
	Lys (7)	2 kDa	TCT	49	0.45	2.7
	Lys (7,8)	5 kDa	TCT	93	0.34	4.7
	Lys (9)	1.1 kDa	OTs	20	0.71	1.2
	Lys (9)	2 kDa	OTs	69	0.66	4.1
	Lys (9)	5 kDa	OTs	68^h	0.48	3.4
					(0.88)^h	(6.1)^h
PHPMA	Lys (n.g.)	7 kDa^d	CDI	14^h	0.14	0.63	Porcine pancreas^80
					(0.12)^h	(4.6)^h
Carbohydrate polymers
Dextran	Lys(n.g.)	40 kDa	CNBr	53^f, 7^g	0.078	—	Multi-site attachment. 11 wt% enzyme^81
					(0.078)^f	—
Dextran	Gln (n.g)	72 kDa	TGase	69^i, 50^g	0.0348	7.5	Multi-site attachment. 1.5, 1.8, and 0.7 chains per enzyme^82
CMC	Gln (n.g)	25 kDa	TGase	82^i, 61^g	0.0150	9.33
PSucr	Gln (n.g)	69 kDa	TGase	56^i, 33^g	0.0322	7.0
					(0.0355)^i	(12.3)^i
Dextrin	Lys (n.g.)	7.7, 39–55 kDa^e	CDI	51–69	—	2.54–4.09	Porcine pancreas.
Dextrin	Lys (n.g.)	47, 48–123 kDa^e	CDI	34–63^h	—	1.25–2.69	Multi-site attachment^79
					—	(2.52–2.93)^h
Dextrin	Lys (n.g.)	2.2, 22 kDa^d	CDI	15	0.13	0.69	Porcine pancreas. Multi-site attachment^80
Dextrin	Lys (n.g.)	36, 51 kDa^d	CDI	19^h	0.11	0.85
					(0.12)^h	(4.6)^h

---

Table 4 Structural and enzymatic properties of lysozyme-polymer conjugates

Polymer	Residue (number)^a	MW(chain, conjugate)^b	Coupling method	Activity (%)	KM/mM (native)	kcat/s^−1 (native)	Comment, ref.
a Number of residues modified on protein. b Molecular weight of conjugate only given for multi-site attachment. Unless identified by superscript c (Mn) the nature of the average molecular weight was not specified. d Glycol chitin (water soluble chitin). e Micrococcus lysodeikticus. f p-Nitrophenyl tetra-N-acetyl-β-chitotetraoside. g Micrococcus Luteus.
P(DL-alanine)	Lys (5.8)	1.9 kDa^c	NCA	63^d, 2^e	—	—	Grafting from^49
	Lys (5.7)	1.5 kDa^c		73^d, 17^e	—	—
	Lys (5.5)	1.2 kDa^c		76^d, 30^e	—	—
	Lys (5.4)	1.0 kDa^c		81^d, 31^e	—	—
	Lys (3.9)	800 Da^c		85^d, 70^e	—	—
	Lys (3.2)	700 Da^c		79^d, 85^e	—	—
MPEG	Asp (1)	550 Da	CDI	90	—	—	Site specific^47
	Asp (1)	2 kDa		75	—	—
	Asp (1)	5 kDa		38^d	—	—
MPEG-deg	Lys (5,6)	5 kDa	NHS	0 → 60^e	—	—	Degradable linker^110
MPEG-deg	Lys (1–6)	5 kDa	NHS	∼0–5 → 100^e	—	—	Degradable linker^111
MPEG-deg	Lys (1)	5 kDa	NHS	∼0→∼50–95	—	—	Several degradable linkers^60
MPEG-deg	Lys (5,6)	5 kDa	NHS	∼0→∼40–90^e	—	—
MPEG1200-PAsp34	Lys (0)	∼5.2 kDa	NC	∼230	—	—	Small-molecule substrate^105
MPEG1200-PAsp34	Lys (3–5)	∼5.2 kDa	GA	∼230^f	—	—
PNIPAAm	Cys (1)	n.g., ∼20–40 kDa	SS and Mal	∼100^e	—	—	V131C mutant. Grafting from^62
MPEG	Lys (1)	12 kDa	NHS	2.3 → 9.6	—	—	Certain conjugates contain cleavable linker^61
MPEG-deg	Lys (1)	12 kDa	NHS	2.8 → 97.4	—	—
MPEG-deg	Lys (1)	20 kDa	NHS	1.4 → 101.6	—	—
MPEG	Lys (4,5)	12 kDa	NHS	1.5 → 2.3	—	—
MPEG-deg	Lys (4,5)	12 kDa	NHS	1.5 → 95.8	—	—
MPEG-deg	Lys (3,4)	20 kDa	NHS	1.1 → 99.7^e	—	—
PVPy-b-PEO10000	Lys (∼1)	20 kDa^c	RA	20–100^e	—	—	Micellar system^106
MPEG	Lys (∼1)	2.3 kDa	NHS	80^e	—	—	Ref. 59
PMPC	Lys (n.g.)	35.6 kDa^c	NHS + SS	∼100^g	—	—	Ionic polymer. After heating^66

---

Table 5 Structural and enzymatic properties of papain-polymer conjugates

Polymer	Residue (number)^a	MW (chain, conjugate)^b	Coupling method	Activity (%)	KM/mM (native)	kcat/s^−1 (native)	Comment, ref.
a Number of residues modified on protein. b Molecular weight of conjugate only given for multi-site attachment. Unless identified by superscript c (Mw) the nature of the average molecular weight was not specified. d N α-Benzoyl-L-arginine-ethyl ester. e N α-Benzoyl-DL-arginine-p-nitroanilide. f Polymer conjugated to BMA block.
CAP	Lys (n.g.)	∼60 kDa, n.g.	CDI	27	—	—	Multi-site attachment. Anionic polymer^97
Eudragit-S	Lys (n.g.)	∼135 kDa, n.g.	CDI	40	—	—
HP-55	Lys (n.g.)	∼45 kDa, n.g.	CDI	17	—	—
MPM-06	Lys (n.g.)	n.g., n.g.	CDI	80^e	—	—
PSucr	Lys (n.g.)	400 kDa, n.g.	RA	103	0.94	0.067	Multi-site attachment. Hyperbranched polymer^83
PSucr	Lys (n.g.)	400 kDa, n.g.	RA	97	1.09	0.059
PSucr	Lys (n.g.)	400 kDa, n.g.	RA	81^e	1.00	0.050
					(1.00)^e	(0.064)^e
PMPC	Lys (7–8)	5 kDa^c	NHS	41.5	—	—	Ionic polymer^96
PMPC	Lys (14–15)	5 kDa^c	NHS	34.1^d	—	—
PMPC	Lys (14–15)	5 kDa^c	NHS	30–35^d	—	—	Ionic polymer^54
PMPC	Lys (12–13)	12 kDa^c	NHS
PMPC	Lys (12)	24 kDa^c	NHS
PMPC	Lys (6–7)	37 kDa^c	NHS
MPEG	Lys (12–13)	5 kDa^c	n.g.
PMPC	Lys (6–7)	11 kDa^c	CDI	41^d	—	—	Amphiphilic polymer^107
P(MPC-ran-BMA5%)	Lys (6–7)	10 kDa^c	CDI	35^d	—	—
P(MPC-ran-BMA25%)	Lys (6–7)	10 kDa^c	CDI	32^d	—	—
P(MPC-ran-BMA50%)	Lys (6–7)	12 kDa^c	CDI	24^d	—	—
P(MPC-b-BM5%)	Lys (6–7)	11 kDa^c	CDI	33^d	—	—
P(MPC-b-BMA25%)	Lys (6–7)	11 kDa^c	CDI	33^d	—	—
P(MPC-b-BMA50%)	Lys (6–7)	10 kDa^c	CDI	12^d	—	—
P(BMA-b-MPC5%)^f	Lys (6–7)	12 kDa^c	CDI	30^d	—	—

---

Table 6 Structural and enzymatic properties of bovine serum albumin-polymer conjugates

Polymer	Residue (number)^a	MW (chain, conjugate)^b	Coupling method	Activity (%)	KM/mM (native)	kcat/s^−1 (native)	Comment, ref.
a Number of residues modified on protein. b Molecular weight of conjugate only given for multi-site attachment. Unless identified by superscript c (Mn) the nature of the average molecular weight was not specified. d 4-Nitrophenylacetate.
PNIPAAm	Cys (1)	148 kDa^c	SS	95^d	—	—	Grafting from^63
PNIPAAm	Cys (1)	234 kDa^c	Mal	95^d	—	—	Grafting from^64
PPEGA	Cys (1)	7.1 kDa^c	VS	92^d	—	—	Partially reduced protein^65
PMPC	Cys (1)	35.6 kDa^c	SS	∼100^d	—	—	Ionic polymer^66
PM13	Lys (18)	13 kDa, n.c.	MA	63^d			Multi-site attachment. Comb-shaped^92
PM100	Lys (12)	100 kDa, n.c.	MA	93^d

---

Table 7 Properties of selected conjugates^a

Polymer	Coupling	Functionality	Activity			Thermal stability
			Degree of conjugation	Length of chains	Altered pH optima
a (SP) single-point attachment; (MP) multi-point attachment; (N) neutral; (HB) hydrogen bonding; (C) charged; (×) little, no, or ambiguous effect; (+) positive effect; (–) negative effect.
ASNase from E. coli
MPEG	SP	N	−^33	−^33
MPEG2	SP	N	−^34,35			×^74
P(DL-alanine)	SP	N	−^48	−^48	no^48	×^48
Silk fibroin	MP	N	^72			+^72
PM13 and PM100	MP	C				+^74
PMPEG5000-g-VP-co-MA)	MP	C			yes^89	−^89
P(VP-co-MA)	MP	C			yes^90	+^90
Sericin	MP	HB/C			yes^93	+^93
N,O-CM-Chit	MP	HB/C			yes^94
ASNase from E. carotovora
MPEG	SP	N	+^37,38
MPEG2	SP	N	+^37,38
levan	MP	HB		–^73	no^73	+^73
αCT
MPEG	SP	N	−^41	×^40
PHPMA	SP	HB	×^50	×,^50 +^51	no^77	+^77
PPEGMA	SP	N	−^53
Dextran	SP/MP	HB	−^55	×^55	no^78
PDMAEMA	SP	C			no^53
P(NIPAAm-co-AADG)	MP	HB				×^75
PAGM	MP	HB				+^76
PNaPS	SP	C			no^53
CMC	MP	HB			no^78
PSucr	MP	HB			no^78
CM-PβCD	MP	HB				+^98
Trypsin
MPEG	SP	N	+^43,45	−,^43,46 +^46
MPEG2	SP	N	+^37,43
PDMAPS					no^95
PNIPAAm	SP	N	+^52
Dextrin	MP	HB				+^80
Dextran	MP	HB			no^81	+^81
Dextran (aminated)	MP	HB/C			yes^82	+^82
CMC (aminated)	MP	HB/C			yes^82	+^82
PSucr (aminated)	MP	HB/C			yes^82	+^82
Lysozyme
MPEG	SP	N		−^47
P(DL-alanine)	SP	N	−^49	−^49	yes^49
Papain
PMPC	SP	C	×^54	×^54
PSucr	MP	HB			no^83	+^83
CAP, Eudragit-S, HP-55, MPM-06	MP	C				+^97

---

2.1. Lessons learned from PEGylation

In all but one recent example, modification of L-asparaginase (ASNase) has been directed towards the amino groups on the protein, as seen in Table 1. In one of the first published examples of PEGylation of Escherichia coli (E. coli) ASNase using cyanuric chloride coupling agent, Ashihara et al.³³ have modified the protein with between 18 and 77 copies of α-methoxy-poly(ethylene glycol) (MPEG) with molecular weights ranging between 750 Da and 5 kDa and have evaluated enzymatic activity using two different substrates. The authors demonstrated that both increased MPEG length and number of copies caused a decrease of enzymatic activity. A similar effect with respect to the number of chains was observed for double branched MPEG₂ (2 × 5 kDa) by Matsushima et al. and Zhang et al. for E. coli ASNase.^34,35 One interesting finding, which came through our analysis of PEGylated ASNase in Table 1, is that conjugates derived from Erwinia carotovora (E. carotovora) maintained higher activities than comparable conjugates prepared from E. coli ASNase. For example, Soares et al.³⁶ have prepared a conjugate from E. coli ASNase bearing 50 copies of MPEG (5 kDa) introduced by N-hydroxysuccinimide (NHS) coupling and which has an activity of 30%, in comparison to an equivalent conjugate prepared by Monfardini et al.³⁷ from E. carotovora and with an activity of 110%. In fact, for E. carotovora ASNase, activity was higher when modified with branched MPEG₂versus linear MPEG.^37,38 Of course, as ASNase obtained from E. coli and E. carotovora are serologically and biochemically different from one another,³⁹ such differences are not surprising. In the remainder of this contribution these two proteins are treated independently. Both Monfardini et al.³⁷ and Veronese et al.³⁸ observed that E. carotovora ASNase coupled with either MPEG or MPEG₂ possessed higher enzymatic activity than the native enzyme. This phenomenon has been observed for other enzymes (as evidenced below) and in the present case was ascribed to the production of a more active form of the enzyme given that the K_M values of the modified forms were not changed upon modification.

Rodríguez-Martínez et al.⁴⁰ have performed a systematic evaluation of the activity of alpha-chymotrypsin (αCT) modified with 1–8 copies of NHS derivatives of MPEG with molecular weights ranging from 700 Da to 5 kDa (Table 2). The authors have found that both k_cat and K_M were affected by PEGylation, though these effects were independent of molecular weight. They have observed a decrease in k_cat with the number of polymer chains attached concurrent with an increase in K_M, which they attribute to an increase of the rigidity of the protein. This effect on K_M, however, appears to be more pronounced for the shorter (≤ 2 kDa) MPEG chains versus 5 KDa chains. Castellanos et al.⁴¹ have also demonstrated that the number of copies of MPEG 5 kDa coupled to αCT correlated with decreased activity. These authors however, also showed a noticeable increase of K_M with number of MPEG (5 kDa) chains attached. This trend was also observed by Chiu et al.⁴² who observed that the MPEG-modified αCT had well-preserved or even elevated k_cat values towards low molecular weight substrates.

Gaertner et al.⁴³ have shown that the activity of trypsin–MPEG conjugates with molecular weights ranging from 350 Da to 10 kDa increased dramatically in comparison to the native protein (Table 3). This phenomenon has been reported by others and has been attributed to a change in the local environment of the enzyme.⁴⁴ Enzymatic activity, while always higher than the native (both esterase and amidase activity) decreased as a function of polymer chain length. The authors also observed a two-fold decrease of K_M, while only a slight effect of MPEG molecular weight on k_cat.⁴³ The authors remarked an increase in activity of trypsin when modified with the branched MPEG₂ (2 × 5 kDa) via the cyanuric chloride method. Abuchowski et al.⁴⁵ have noted an increase in esterolytic activity of trypsin by increasing the number of chains of MPEG on the protein from 3 to 8–9, though a decrease of activity towards peptide and protein substrates. Monfardini et al.³⁷ have also observed this increase of esterolytic activity upon attachment of MPEG₂. Treetharnmathurot et al.⁴⁶ have measured the relative activity and kinetic parameters for MPEG–trypsin conjugates prepared by three different coupling methods. All these conjugates had lower activity than the native trypsin. Within the conjugates prepared, the authors appeared to obtain dissimilar trends as a function of molecular weight between 1.1 and 5 kDa for the three different coupling strategies. That is, activity increased as a function of molecular weight when coupled using cyanuric chloride or a tosylated MPEG, but decreased when carbodiimide conjugation was used for coupling. This phenomenon is discussed in Section 3.1.

So et al.⁴⁷ have observed a decrease in enzymatic activity of lysozyme–MPEG conjugates as a function of polymer molecular weight from 550 Da to 5 kDa (Table 4). The polymer was coupled in a site-specific manner to the aspartic acid residue at position 119 on the protein. This residue is distant from the protein's active catalytic site. The authors rationalize their findings by spreading of the MPEG chain over the surface of the protein resulting in interference with the protein's active site cleft.

2.2. Other polymers

Poly(DL-alanine). In Tables 1 and 4 poly(DL-alanine) has been grown from both E. coli and E. carotovora ASNase as well as from lysozyme. Uren and Ragin⁴⁸ observed no change in the K_M of the modified ASNases, and only slight changes in the pH optima and temperature dependent enzymatic activity. In general, the authors indicated that enzymatic activity decreased as a function of extent of conjugation. For the “grafting from” approach used to prepare these conjugates (Section 3.3) both the number and length of the polymer chains vary simultaneously and thus the individual effects of these cannot be distinguished. For the modified lysozyme, Yoshimura et al.⁴⁹ observed that the enzymatic activity of the conjugate towards glycol chitin decreased proportionally with the average length of the polyalanine chain, which the authors have attributed to a steric hindrance effect. The loss in activity towards Micrococcus lysodeikticus (M. lysodeikticus) was greater than that toward glycol chitin. The authors indicate that the lytic activity of the conjugate against M. lysodeikticus appeared to depend on the number of modified amino groups (when less than five).

Poly(N-(2-hydroxypropyl)methacrylamide) (PHPMA). Lu et al.⁵⁰ have modified αCT with between 3 and 5 copies of semi-telechelic PHPMA with molecular weights ranging from 2.7 to 5.5 kDa. In addition, the authors have prepared a macromolecular substrate composed of a PHPMA backbone bearing peptide p-nitroanilide side chains. The ability of the conjugates to catalyze the hydrolysis of the polymer-bound substrate was similar or lower than the native enzyme, with small variations. The authors notably showed that the degree of conjugation and polymer molecular weight did not have a pronounced effect on enzymatic activity towards this substrate (Table 2). Oupický et al.⁵¹ have also modified αCT with semi-telechelic PHPMA and have observed that catalytic activity towards a small-molecule substrate increased as a function of polymer molecular weight from 2.7 to 10.9 kDa. However, the authors have observed a significant decrease in proteolytic activity towards a PEGylated substrate and towards bovine serum albumin (BSA).

Poly(N-isopropylacrylamide) (PNIPAAm). Ding et al.⁵² have conjugated PNIPAAm to trypsin and have observed a remarkable increase of both esterolytic and amidase activity with the number of conjugated polymer chains (Table 3). This phenomenon was rationalized to result from a change in microenvironment of the enzyme active site as will be discussed in Section 6.

Poly(polyethyleneglycol monomethyl ether methacrylate) (PPEGMA). Lele et al.⁵³ have modified αCT with α-bromo amide initiating groups and have conducted aqueous atom transfer radical polymerization (ATRP) of PEGMA (1.1 kDa and 475 Da side-chains) from the modified protein. The authors observed that conjugate activity decreased as the number of conjugated PPEGMA chains increased (Table 2). We have unfortunately not found examples of conjugates prepared with polyethylene glycol methacrylate (i.e., bearing a hydroxyl versus an ether end group) and which fit in to the scope of this review.

Poly(2-methacryloyloxyethyl phosphorylcholine) (PMPC). Miyamoto et al.⁵⁴ have modified papain with PMPC and the resulting conjugates possessed lower activity than the native protein (Table 5). However, the authors have observed no correlation between residual enzymatic activity and either the degree of conjugation from 6–15 chains or polymer molecular weight from 5 to 37 kDa.

Dextran (via single-point attachment). Solà and Griebenow⁵⁵ have modified αCT with mono-(dextranamido)-mono-(succinimidyl) suberate (10 kDa), a dextran bearing a single reactive site for reaction with the protein. For the resulting conjugates, the authors observed that k_cat was reduced as a function of the polymer molar content, independently of the polymer's molecular weight. The K_M of the conjugates remained unchanged (Table 2).

2.3. General trends

Based on the findings of the previous two sections, it becomes obvious that dissimilar trends between enzymatic activity and the number/length of conjugated polymer chains are observed for the selected proteins within this contribution (Table 7). It is interesting to remark that certain proteins such as E. coli ASNase and E. carotovora ASNase display opposite trends with respect to the number and length of conjugated polymer chains, despite their related structure and function. This is an indication that the enzymatic properties of proteins are highly sensitive to polymer modification as the latter cannot only act as a diffusional barrier for the substrate, but can also modify the three-dimensional structure of the protein, thus rendering it either a more or less efficient enzyme. This also means that predicting the effect of polymer modification on the enzymatic activity of a particular enzyme is not trivial.

In general, within the dataset examined, all vinyl polymers obeyed the same trend as PEGylation (albeit to a different extent) for each individual protein. This is a first indication that polymer-modification per se has an influence on protein activity. However, as highlighted by Treetharnmathurot et al.,⁴⁶ coupling chemistry may also play a role on activity, notably because the authors have observed different trends with respect to molecular weight for different coupling methods (Table 7).

3. Polymer coupling strategy

With the currently available (bio)chemical toolbox for preparing protein–polymer conjugates, there are many possible ways of preparing such systems which go well beyond the random convergent coupling of a semi-telechelic polymer to a protein. The polymer coupling strategy selected for preparing the protein–polymer conjugate has a strong potential to affect the properties of the protein. This is because functional groups of diverse nature are directly added to the protein, and because these may be added in locations important for e.g. catalytic activity. Moreover, the preparative methods used to synthesize/purify the conjugates prepared by the divergent “grafting from” approach as well as the multiplicity of the bonds between a protein and a polymer can also play a key role in conjugate properties. In this section, we examine the effect of these four parameters on conjugate bioactivity. In each subsection we attempt to draw general conclusions from the available data.

3.1. Ligation chemistry in the “grafting onto” approach

Within the dataset examined, there are only two studies in which several ligation chemistries are used to conjugate the terminus of a semi-telechelic polymer to a protein. Treetharnmathurot et al.⁴⁶ have measured the relative activity and kinetic parameters for MPEG–trypsin conjugates prepared by three different coupling methods. All resulting conjugates had lower activities than the native trypsin. As stated previously, the authors appeared to obtain dissimilar trends as a function of molecular weight for the three different coupling strategies. That is, activity increased as a function of molecular weight when MPEG was coupled either using cyanuric chloride or a tosylated form of the polymer, but decreased when a carbodiimide was used to couple a succinoylated MPEG to the protein. Concurrently, k_cat decreased as a function of molecular weight for both alkylating coupling chemistries (i.e., cyanuric chloride and tosylate derivative) but increased for the acylating coupling chemistry (carbodiimide). The K_M of all conjugates decreased as a function of molecular weight. From this study it appears as though, for trypsin, ligation chemistry which maintains the charge of the ε-amino groups of the lysine residues on the protein (albeit with a potential modification of their pK_a), has a positive effect on enzymatic activity. Lu et al.⁵⁰ have modified αCT with semi-telechelic PHPMA either at lysine residues or at aspartic/glutamic acid residues (Table 2). Conjugates prepared by modification of the amino groups on αCT showed higher activities than the native enzyme. However, the authors have observed a decrease of enzymatic activity for equivalent conjugates for which the PHPMA chains are conjugated to carboxylic acid groups on the protein.⁵⁰ It is known that a carboxylic acid group of an aspartic acid is involved in the active site of αCT.⁵⁶ With the limited number of systematic studies available to discuss the effect of conjugation chemistry, it is impossible to draw more general conclusions at this time.

3.2. Random versus site-specific coupling

Site-specificity is an interesting and modern approach for preparing conjugates with potentially less loss of enzymatic activity.³⁰ In this approach, detailed knowledge of the protein's three-dimensional structure is required in order to select locations for polymer-modification far from its catalytic site. Alternatively, one can select a location near or within an antigenic region of the protein in order for the conjugate not to be recognized by the immune system. In this section, we highlight the benefits or disadvantages of site-specific protein modification in comparison to random modification. Of course, as such modifications are specific to a given protein, the latter are treated individually.

ASNase. ASNase from E. coli is a protein, which consists of four identical subunits, each containing a single disulfide bond on the outer edge of the protein, distant from the protein's active site.⁵⁷ Balan et al.⁵⁸ have modified the tetrameric ANSase with exactly four MPEG chains by opening these disulfide bonds and alkylating the cysteine residues (Fig. 1). The authors observe that enzymatic activity of the conjugates was similar to that of the native protein, irrespective of the length of the MPEG chain. These results contrast with those obtained by conventional random PEGylation of E. coli ASNase, as seen in Tables 1 and 7 and as discussed above. The authors rationalize their findings by the distance separating the polymer from the enzyme's active site. However, it was also observed that the conjugates also retained their antigenicity, given that the conjugation site was also distant to the enzyme's antigenic site.

Fig. 1 Tetrameric L-asparaginase from E. coli (DPB entry 4ECA). Thr-12 in active region of each monomer acetylated with L-aspartic acid (magenta). Site-specific introduction of the polymer chains at Cys⁷⁷–Cys¹⁰⁵ (blue) yielded a conjugate with full retention of activity, but inefficient elimination of antigenicity.⁵⁸ Protein image produced with PyMol v.1.1.

Lysozyme. So et al.⁴⁷ have site-selectively modified lysozyme at the aspartic acid residue located at position 119 with MPEG in a multi-step manner, by prior modification of the protein with (2-pyridyl)-dithioethylamine, purification of the modified protein by ion-exchange chromatography to obtain the site-selectively modified protein, reduction of the introduced disulfide bond, and alkylation of the resulting thiol group with a brominated MPEG derivative. Aspartic acid 119 is separated from the active cleft region of the protein (Fig. 2a), though as the authors have observed a decrease of enzymatic activity towards glycol chitin with respect to MPEG molecular weight they rationalized that the polymer molecule nevertheless partially disturbed interaction of the substrate with the active site cleft. For comparison, Higashi et al.⁵⁹ have modified lysozyme in a random fashion at lysine residues with ∼1 chain of 2.3 kDa MPEG and obtain similar enzymatic activity (80%) to the comparable conjugate prepared above (1 chain, 2 kDa, 75% activity),⁴⁷ albeit using a different enzymatic substrate. For MPEG chains of 5 kDa or higher, conjugation of even a single polymer chain to lysine fully deactivates the enzyme as seen by several authors in Table 4.^60,61 In contrast, 38% enzymatic activity (towards glycol chitin) is retained when the 5 kDa MPEG chain is uniquely located at aspartic acid-119, clearly demonstrating in this instance the benefit of site-specific protein modification.⁴⁷ The authors attribute this phenomenon to the polymer not spreading along the surface and wrapping around the enzyme but rather adopting a compact conformation around the conjugation site. The authors also observed that site-specific PEGylation (Asp-119) did not reduce T cell activation in comparison to native lysozyme. Heredia et al.⁶² have site-specifically mutated lysozyme to introduce a cysteine residue in lieu of the valine residue at position 131 (V131C, Fig. 2b). Initiators for ATRP were then introduced at this location either via disulfide exchange or by Michael addition and exploited for the polymerization of NIPAAm. The authors have selected this location for modification because of its distance from the enzyme's active cleft. They observed that conjugation with the polymerization initiator or with the polymer at this location did not decrease enzyme activity. In fact, catalytic activity was maintained despite the polymer having a molecular weight of 20–40 kDa.

Fig. 2 (a) Hen egg lysozyme (PDB entry 2HU1) with residues implicated in catalysis (Glu-35 and Asp-52) in magenta and Asp-119 in blue. (b) T4 lysozyme V131C (PDB entry 2HUK) with residues implicated in catalysis (Glu-11 and Asp-20) in magenta and Cys-131 in blue. For both proteins, lysine residues are in red. Conjugation at Cys-131 lead to full retention of enzymatic activity (antigenicity not tested). Modification at Asp-119 lead to partial loss of activity and did not eliminate antigenicity.^47,62 Protein images produced with PyMol v.1.1.

Bovine serum albumin. BSA has been modified with PNIPAAm, poly(polyethylene glycol monomethyl ether acrylate) (PPEGA), and PMPC in a site-specific manner at the sole free reduced cysteine residue at position 34 with seemingly full retention of esterolytic activity (Table 6).^63–66 Interestingly, Córdova et al.⁶⁷ have very recently shown that BSA retains complete catalytic activity at high temperatures (up to 160 °C), even in the presence of 25 mM sodium dodecyl sulfate. This indicates that a catalytically competent orientation of amino acid residues exists in the denatured or partially unfolded protein, though disruption of the protein disulfide bonds reduced esterase activity to ∼50%. When De et al.⁶⁴ have prepared a BSA conjugate with PNIPAAm, they observed that the latter maintained 92% of its activity at 25 °C, though above the LCST the activity was reduced to 75%. These two phenomena may be related, though additional investigation is required. In light of the findings for this protein, it is perhaps not unambiguous to correlate maintenance of protein structure or site-specific modification to enzymatic activity.

Overall the examples seen in this section demonstrate that the location chosen for protein modification is crucial for obtaining a conjugate with desired properties. Indeed, when the location is properly chosen, the characteristics of the polymer, within the dataset examined, appear to play a secondary role on bioactivity.

3.3. “Grafting from”

The so-called “grafting from” approach for preparing protein–polymer conjugates by direct polymerization from a protein is associated with many advantages, including: the possibility of accurately characterizing the sites of polymer modification, the ease of purification of the conjugate, and the greater uniformity of the produced conjugates.^18,23,27 However, in order for the “grafting from” approach to be viable, the protein must survive modification with the initiating moiety, the polymerization reaction (which can involve transition metals or ionizing radiation), and the purification procedure without compromising significantly activity. In this section, we discuss aspects of the synthesis, the purification, and the opportunities offered by this approach, in particular in as much as how they affect enzymatic activity.

Organic (co)solvents. Uren and Ragin⁴⁸ have prepared conjugates of ASNases with poly(DL-alanine) by direct polymerization of DL-alanine-N-carboxyanhydride from the protein. One interesting feature of this reaction is that the total number of amino groups on the conjugate remains the same as that of the native protein, given that the consumed amino group on the initiating lysine residue is replaced with the N-terminus of the grafted polypeptide chain. However, owing to different solubility characteristics of the protein and the monomer, the authors must perform polymerization in 1 : 1 dimethyl sulfoxide : aqueous buffer. Depending on the desired characteristics of the conjugate, either 100 mM K₂HPO₄ (pH 6.8, 0 °C) or 50 mM NaHCO₃ (pH 8.6, 4 °C) was used. Yoshimura et al.⁴⁹ have encountered a similar problem for the preparation of conjugates of lysozyme with the same polymer. In this case, the authors performed their polymerizations in 20 vol% dioxane in 50 mM phosphate buffer (pH 7.1, 0 °C). In both previous examples, the proteins maintained reasonable enzymatic activity following exposure to organic solvents. In particular, conjugates prepared with lysozyme maintained activity towards glycol chitin to a similar (or even better) extent to conjugates prepared with MPEG (Table 4) by other authors. This is an indication that, to a certain extent, organic solvents are acceptable for improving the homogeneity of the polymerization medium without drastically affecting the enzymatic activity of the conjugate.

Modification with initiators and exposure to electromagnetic irradiation. Ito et al.⁶⁸ have prepared conjugates of trypsin with either the negatively charged pH-sensitive copolymer of methyl methacrylate and methacrylic acid or the positively charged redox-sensitive poly(3-carbamoyl-1-(p-vinylbenzyl)pyridinium chloride). In this investigation, about 6–7 azo groups were connected to amino groups in trypsin in aqueous buffer at 4 °C, the modified protein purified by dialysis, and then used to initiate graft polymerization of the appropriate monomers under UV irradiation (120 W mercury lamp, 24 h). The catalytic activity of azo-containing trypsin decreased to about half that of the original enzyme though it is unclear whether this is the result of the introduction of these groups or due to an uncharacterized degradation process concurrent with the purification procedure (the modified enzyme, which is a protease, was dialyzed for 2 days). Following polymerization, the conjugates were precipitated and purified by size-exclusion chromatography. The resulting conjugates maintained about 16% of the activity of the native trypsin. While the authors ascribe this decrease to conformational changes induced by the graft polymerization or to coverage of the catalytic site, it is also conceivable that the UV-irradiation used during the polymerization contributed to deactivation. Along this line, Liu et al.⁶⁹ have evaluated the effect of γ-irradiation on BSA as a control experiment for the γ-irradiation-initiated reversible addition-fragmentation chain transfer (RAFT) polymerization of PEGA from a BSA macro-RAFT agent. The authors showed that the enzyme retained 92% of its original activity after a six hour exposure, indicating that while major damage to the protein did not take place, partial deactivation following an unknown mechanism is possible. These data indicate that certain polymerization conditions may indeed lead to deactivation of the enzyme and appropriate control experiments are required to account for this.

Purification procedure. Heredia et al.⁶² have prepared a PNIPAAm–lysozyme conjugate through site-selective modification of a genetically introduced cysteine residue with an ATRP initiator. The polymerization from lysozyme was conducted in water at ambient temperature in the presence of resin-bound sacrificial initiator. Following removal of the resin, the solution was warmed to precipitate the conjugate, which was then easily collected. While, ideally, after this stage pure conjugate should be obtained, the authors observe that non-unitary initiator efficiency from the protein resulted in a mixture of conjugate and native protein, which were subsequently separated by preparative size-exclusion chromatography. Following this procedure, purified conjugates with enzymatic activity equal to that of the native enzyme were obtained. This demonstrates that this specific purification procedure is not adversely affecting protein structure or function. In a similar example, De et al.⁶⁴ have prepared PNIPAAm–BSA conjugates by RAFT polymerization from a macro-RAFT agent prepared by modification of the BSA at its single reduced cysteine residue. This was accomplished in aqueous media (pH 6, 26 °C) and isolation of the conjugate was achieved by thermal precipitation of the responsive conjugate at 40 °C. Though, as for lysozyme, residual BSA remained following this procedure. In general these studies illustrate that within the “grafting from” approach it is possible to purify the resulting conjugate with little adverse effect on protein activity. From these examples, it appears as though until quantitative initiator efficiency is achieved, it is not possible to directly prepare protein–polymer conjugates bearing a single polymer chain by the “grafting from” approach that are devoid of non-modified protein contaminant. The latter must necessarily be removed in a subsequent purification step.

Overall, the examples discussed in this section demonstrate that it is possible to maintain high levels of protein bioactivity following synthesis and purification of conjugates prepared by the “grafting from” approach.

3.4. Multi-point covalent bonding (reactive polymer side-chains)

In the previous sections, the modification of proteins with semi-telechelic polymers has been discussed. However, the multi-point attachment of enzymes to polymers (i.e., the polymer bears multiple protein-reactive groups along its backbone) has been widely studied and is considered to be the most general approach to stabilize these proteins against different denaturing conditions,⁷⁰ in particular for avoiding their inactivation caused by protein unfolding. This holds particularly true for enzymes composed of multiple subunits, such as ASNase, and whose enzymatic activity is lost due to dissociation into inactive subunits through dilution.⁷¹ Consequently, and as highlighted in this section, multi-point attachment has the potential to increase the thermal stability of the conjugate because of its preventative effect on protein denaturation as well as to affect the pH-activity profiles of the conjugate because of the change in the microenvironment surrounding the enzyme.

In the multi-point attachment approach, because both the polymer and the protein possess multiple functional groups susceptible to react together, the resulting conjugates are, in general, less well-defined than their analogs prepared by single-point attachment. One ambiguity which must be clarified is that, in contrast to conjugates prepared by single-point attachment, the degree of modification of the reactive residues on the protein is not necessarily proportional to the number of conjugated polymer chains. As a consequence, for these conjugates the molecular weight of the final conjugate is a second necessary parameter for describing the conjugate. Unfortunately, for many of the conjugates listed in Tables 1–6, this important parameter is not defined and thus analysis of the influence of polymer attachment is somewhat hampered. In this section we discuss the effect of multiple covalent coupling of proteins to polymers and do not discuss any specific non-covalent interactions between the two. We have, however, included in this section polymers bearing hydroxyl groups, which have the potential to participate in hydrogen bonding with the protein, because we found it was not possible to distinguish the effect of this type of non-covalent interaction from that of multiple covalent coupling.

ASNase. Modification of ASNase by multi-point attachment to silk fibroin with gluteraldehyde was performed by Zhang et al.⁷² Addition of enzyme substrate was required to protect the active site from the cross-linking reaction. Modification with silk fibroin resulted in an increased optimal enzymatic reaction temperature, a widened optimal reaction temperature range, and an increased stability towards tryptic digestion (Table 7). Enzymatic activity appeared to be relatively insensitive to the degree of modification of amino groups over a broad range for this protein, though decreased at very high values. Gluteraldehyde concentration affected significantly the activity of the enzyme, which was found to increase proportionately with gluteraldehyde concentration, then decrease after a maximal value. This has been attributed to the opposing effects of protein stabilization via conjugation and reduction of activity upon cross-linking. The half-life of the modified ASNase lengthened to nearly 84 h, about twice that of the native enzyme, suggesting that covalent conjugation also enhanced the resistance to proteases in human serum. Vīna et al.⁷³ have coupled oxidized levan (a highly branched fructose polysaccharide from Zymomonas mobilis) to E. carotovora ASNase by multi-point attachment via reductive alkylation and have demonstrated that the loss of ASNase activity due to conjugation was proportional to the molecular weight and amount of coupled levan. This decrease may be explained by the inability of the enzyme and substrate to interact due to the increasing steric hindrance imparted by the branched levans. The apparent K_M values were 1.5–4 times higher than that of the native enzyme, a finding which the authors rationalize to not result from a change in ASNase itself, but rather due to the diffusional barrier imposed by the branched polysaccharide. The stability of the conjugates towards thermal denaturation was improved in comparison to the native enzyme, an effect which is rationalized by the authors as resulting from greater rigidity of the enzyme and increase activation energy for “uncoiling” due to the multi-point covalent attachment of the polymer. The conjugates possessed a broadened optimum pH-activity range, without a significant shift in its maximum in comparison to the native protein. Kodera et al.⁷⁴ have modified ASNase with two comb-type polymers, the first being a copolymer of poly(oxyethylene) allyl methyl diether and maleic anhydride (PM₁₃) with MW = 13 kDa and the second a copolymer of poly(oxyethylene) 2-methyl-2-propenyl methyl diether and maleic anhydride (PM₁₀₀) with MW = 100 kDa (Fig. 3a). The two polymers were conjugated to ASNase by reaction of the maleic anhydride moieties along the backbone with the amino groups on the protein. The authors compared the properties of these conjugates to those prepared with linear and MPEG₂ (2 × 5 kDa, coupled via TCT chemistry). The effect of modification towards heat, urea, and acidity were tested. The authors have found that while native ASNase lost enzymatic activity within 30 min at 65 °C, the enzymes modified with the anhydride-bearing polymers retained between 35 and 90% of their activity. Similarly, PM₁₃ and PM₁₀₀-modified ASNase maintained between 70 and 85% of their initial activity in 4 M urea, conditions under which the native enzyme lost activity in 10 min. Also, at pH 4, the native enzyme maintained 10% of its original activity after 1 h incubation, while the polymer-conjugated enzymes retained ∼80% of their activity. The MPEG₂ modified ASNase did not show stabilizing effects towards temperature or acidity, which the authors ascribe to the single reactive site on these polymers (Table 7). The authors also show that the stabilizing effect is dependent upon polymer molecular weight. The influence of backbone architecture of these comb-type polymers on enzymatic activity is discussed in Section 4.3.

Fig. 3 (a) Comb-type polymers developed by Inada and co-workers for multi-point attachment to ASNase and BSA.^74,92 (b) Human serum albumin (PDB entry 1AO6) with residues implicated in esterase activity in (Asn-391, Arg-410, and Tyr-411) magenta, and Cys-34 in blue. Several semi-telechelic polymers have been site-selectively conjugated to Cys-34 of bovine serum albumin (pdb structure not available) with excellent retention of activity. Multi-point attachment of comb-type polymers PM₁₃ and PM₁₀₀ to lysine residues (red) lead to a moderate decrease in activity. Protein image produced with PyMol v.1.1.

αCT. To illustrate the importance of reaction conditions on the structure of the conjugate in the multi-point attachment approach, Kim and Park⁷⁵ have prepared a poly(N-isopropylacrylamide-co-acrylamido-2-deoxy-D-glucose) (PNIPAAm-co-AADG) copolymer, which contains pendant glucose residues, and, upon activation, is conjugated to αCT by reductive alkylation. The authors found that slow addition of the copolymer solution to the protein solution is critical for achieving multi-site coupling of αCT to a single copolymer chain or several copolymer chains. In contrast, other conditions favor gelation due to the formation of extensively cross-linked enzyme–polymer adducts. In fact, the relative molecular weights of the polymer and enzyme as well as the weight percentages of enzyme within the conjugate (14.7–29.4 wt%) suggest that multiple enzymes are conjugated to a single polymer chain. Enzyme activity of the conjugates decreased with increasing proportion of carbohydrate monomer within the feed. Interestingly, this parameter, however, had little effect on the degree of modification of the protein indicating either a change in the properties of the polymer or the implication of non-covalent interactions such as hydrogen bonding on enzyme activity. K_M values of the conjugated enzymes slightly increased as compared with that of native enzyme, though, interestingly, the conjugated enzymes with increasing glucose-monomer content did not show any significant improvements in thermal stability at 40 °C or enhanced activity in organic solvents, compared with the free enzyme. These observations are in contrast with the previous findings by Wang et al.⁷⁶ that αCT conjugated to the carbohydrate moiety (glucose) in a polymethacrylate backbone exhibited robust stability and is also surprising considering the high level of lysine modification of the protein. The authors suggest that the inclusion of NIPAAm in the polymer backbone renders the polymer less hydrophobic than the homopolymer above,⁷⁶ thus increasing the freedom for conformational changes and reducing its efficiency in minimizing thermal unfolding. Unfortunately numerical values for enzyme activity are not available, though k_cat and K_M are stated to be similar to those of the native enzyme. It is also possible that, as the glucose-based homopolymer prepared by Wang et al.⁷⁶ possessed a greater proportion of hydroxyl groups, these may participate in non-covalent interactions with the protein, thus stabilizing it. Lääne et al.⁷⁷ have modified αCT with PHPMA by pre-treatment of the polymer with either cyanuric chloride or hydrazine/NaNO₂, or by introducing active ester moieties along the side-chain by copolymerization of HPMA with a p-nitrophenyl ester of methacryloylated glycylglycine. For all three conjugates the enzymatic activity was quite low, though when related to the content of enzyme within the latter (i.e. the specific activity of the enzyme within the conjugate), the authors indicate that enzymatic activity of αCT is unchanged in comparison to the native form. Thermoinactivation experiments at 50 °C and either pH 6.1 or 8.0 demonstrated improved stability of the conjugates in comparison to the native form of the enzyme. The optimum pH of activity was not modified by conjugation to PHPMA. Sundaram and Venkatesh⁷⁸ have covalently conjugated αCT to various carbohydrate polymers which resulted in reduced specific enzymatic activity depending on the polymer used and the degree of modification of the enzyme. The pH optimum of 8 for the native enzyme was virtually unchanged upon modification (Table 7).

Trypsin. Duncan et al.⁷⁹ have modified trypsin by multi-point attachment to dextrin and have observed a decrease of enzyme activity in relation to an increase in the degree of polymer activation (i.e., for reaction with the protein). In contrast, for those conjugates synthesized using the lower molecular weight dextrin, the degree of activation did not correlate with retained enzyme activity. Calculation of the rate constants K_M, V_max (maximum rate of enzymatic reaction), and k_cat showed that conjugation of trypsin to dextrin typically did not alter K_M, but the V_max was reduced ∼10-fold. The highest degree of trypsin inactivation was achieved using the higher molecular weight dextrin due to the greater steric hindrance caused by the larger polymer chain. Treetharnmathurot et al.⁸⁰ have also modified trypsin with dextrins and, consistent with the previous example, obtained K_M values similar to that of native trypsin. The substrate turnover rate k_cat was decreased 5–7 fold. All the polymer–trypsin conjugates synthesized were less susceptible to autolysis at 40 °C than the native trypsin. Marshall and Rabinowitz⁸¹ have coupled neutral CNBr-activated dextran (40 kDa) to trypsin resulting in a conjugate which showed greater resistance to heat, auto-digestion, and denaturing agents. The enzyme retained significant esterase activity but highly reduced caseinolytic activity compared to the native enzyme. In addition, the resulting conjugates had similar pH-activity profiles to the native trypsin. In another example, Villalonga et al.⁸² have covalently modified the glutamine residues on trypsin with 1,4-diaminobutane-derivatized dextran, carboxymethyl cellulose (CMC), and polymeric sucrose by means of the enzyme transglutaminase (TGase). The thermal stabilization was ascribed to the co-operative contribution of both covalent coupling and the formation of new hydrogen bonds between the covalently linked polysaccharide and the hydrophilic groups at the surface of the enzyme. These attached polysaccharide molecules also prevented protein aggregation at high temperatures, which play an important role in the thermal inactivation mechanism of trypsin.

Papain. Rajalakshmi and Sundaram⁸³ have modified papain with polymeric sucrose at different ratios. The derivatives retained > 80% intrinsic catalytic activity with no change in pH optima and kinetic constants, indicating that the gross tertiary structure was not altered by modification. However, they displayed better thermal stability than native papain, as indicated by their temperature optima being shifted by 10 °C.

Overall, it can be seen in this section that careful choice of synthetic conditions is required for obtaining a reasonably well-defined conjugate. Multi-point attachment of polymers to proteins is an effective means of improving their thermal stability. In the preceding examples, little mention is made of the individual contribution of covalent bonding versus hydrogen bonding on enzyme properties. However, as the polymers in this section are neutral, little or no effect on pH-dependant activity profiles was observed (Table 7). Also, it becomes apparent that the degree of covalent coupling alone cannot account for all observed trends, indicating that polymer properties (such as polarity, architecture, etc.) and the susceptibility of the latter to interact with the protein/substrate play key roles in the enzymatic properties of the conjugate. These will be discussed in turn in the following two sections.

4. Polymer architecture

In this section we survey the influence of polymer backbone architecture on the catalytic properties of protein–polymer conjugates. It is generally known that branched or folded polymers adopt more compact structures than linear polymers in a random-coil configuration. Therefore these polymers may possess a smaller “footprint” on the protein's surface and potentially confer different properties to a conjugate than a flexible linear polymer, which can spread on the protein's surface. Consequently, the way they perform as a diffusion barrier to substrate is likely to be different from that of linear polymers. In this section, we will examine the influence of polymer backbone architecture or propensity to adopt a distinct conformation on conjugate enzymatic properties. We will sequentially discuss polysaccharides which may be linear or hyperbranched, polypeptides with folded structures, and finally comb-type synthetic polymers. Of worthy mention, while MPEG₂ constitutes an elementary branched polymer (from the perspective of the enzyme), the lack of studies which systematically evaluate the length of the polymer segments, the number of conjugated MPEG₂ chains, or the degree of branching (i.e., higher than 2 through an alternative coupling strategy to TCT) unfortunately limits what can be said about the effect of this type of branched polymer on conjugate bioactivity. Comparisons of MPEG₂ to MPEG have already been made in Section 2.1. and will not be treated further.

4.1. Polysaccharides

Polysaccharides have the potential to display different degrees of branching, depending on their source. For instance, CMC is a purely linear polymer, while dextran and dextrin possess a certain non-negligible degree of branching, and bacterial levan and polymeric sucrose are highly branched and adopt an almost spherical structure in solution.^84,85 For a polymer of a given molecular weight, hyperbranched macromolecules adopt more compact and rigid structures in solution than flexible linear polymers. In addition, hyperbranched polymers do not interpenetrate.⁸⁶ Therefore, for an enzyme modified with several polymer chains, these are less likely to interpenetrate to form a uniform diffusion barrier to small molecule substrates, in comparison to a polymer adopting a loose random coil configuration. Clearly, architecture-dependent properties are expected between these classes of polymers. Unfortunately, very little or no information is given in the studies reviewed herein on the degree of branching of these polymers, owing to the difficulties in characterizing this aspect of natural polymers. As such, useful comparisons of the conjugates are difficult to make.

Sundaram and Venkatesh⁷⁸ have evaluated the thermal stability of native and variously modified αCT with polysaccharides including polymeric sucrose (70 and 400 kDa), CMC (12 kDa), and dextran (73 and 250 kDa). The difference in thermal stability is rather unusual in that it decreased from the highly branched polymeric sucrose, to the linear CMC, to the branched dextran. The authors demonstrated that polymer size alone is not the influential factor in improving the stability of a protein, its structure obviously plays an important role, as confirmed by their finding that polymeric sucrose (70 kDa) and dextran (73 kDa) stabilized αCT to different extents though they have about the same molecular mass. Unfortunately, as the molecular weights of the conjugates have not been evaluated, it is not possible to draw more detailed conclusions from this study. The authors rationalized their findings in that the flat ribbon-like chains of CMC would be the most rigid of the three polymers used, based on the suggested interpretation that bulky groups in carbohydrates equatorial to the glycosidic linkage make them so. Additionally, the rigidity of the highly branched polymeric sucrose may also contribute to the enhanced thermal stability of the conjugate. From the measured activity of the conjugates presented in Table 2, the conjugates prepared with a comparable degree of modification of αCT (11–13 and 13–14 lysine residues for CMC and dextran, respectively) only show a small decrease in activity when going from 12 kDa CMC to 250 kDa dextran. In comparison, attachment of a single 5 kDa MPEG chain to αCT by Lele et al.⁵³ led to a comparable degree of inactivation. Also of note is the fact that the conjugate prepared with polymeric sucrose (8–11 reacted lysine residues, 70 kDa per chain) had the same activity as the conjugated prepared with CMC above. Furthermore, the conjugate prepared with 400 kDa polymeric sucrose had higher activity than the conjugate prepared with 200 kDa dextran, indicating that in this case, dextran was more effective than polymeric sucrose in blocking the enzyme's active site. Additional characterization of these conjugates would be necessary to unambiguously establish the reason for this.

In a comparable study, Villalonga et al.⁸² have modified trypsin with dextran (72 kDa), CMC (25 kDa), and polymeric sucrose (69 kDa) by means of the enzyme transglutaminase (TGase). For this coupling reaction, the polysaccharides were oxidized and amino groups introduced with diaminobutane/NaBH₄. These amino groups act as acyl acceptors for the TGase-mediated reaction with γ-carboxyamide groups on the protein. They have noticed that lower thermal stability was conferred to the enzyme after glycosidation with CMC, in comparison with the dextran and polymeric sucrose complexes. No clear differences were observed between the low to moderately branched dextran and highly branched polymeric sucrose. However, the conjugate prepared with polymeric sucrose had a lower proteolytic activity in comparison to the dextran conjugate, despite a lower level of protein modification. In contrast, the esterolytic activity appeared to be less affected. This is in contradiction with the results from the preceding paragraph and points to the need for better characterization of conjugates in order to draw meaningful conclusions for the effect of polymer architecture. The lower thermal stability of the conjugate prepared with CMC was rationalized to result from the conformation of CMC, which adopts a flat ribbon-like structure in solution, thereby causing less interactions (and possibly lower extents of multi-point attachment) in comparison to the more flexible branched dextran or the more-or-less spherical polymeric sucrose.

4.2. Polypeptides

Polysaccharides are not the only polymers to display specific backbone configurations. Indeed, proteins are (in general) linear polypeptides which can adopt compact folded structures with little potential for interpenetration. There is only one example, within the dataset examined, of a conjugate prepared with a folded polypeptide. Poznansky et al.⁸⁷ have modified E. coli ASNase with rat albumin via a multi-point attachment approach using gluteraldehyde, and have managed to maintain activity to 60% of the native protein, despite the presence of an average of 12 large (∼65 kDa) albumin molecules per enzyme (Table 1). However, performing the modification reaction in absence of the substrate L-asparagine resulted in the loss of at least 90% of the enzyme activity. The authors indicated little change in V_max, a 2- to 3-fold increase of K_M, increased thermal stability, and resistance to proteolytic digestion. It is possible that, in contrast to common synthetic polymers, which adopt random coil conformations in solution, the grafted albumin molecules maintain their compact folded structure which would prevent the approach of large molecules such as antibodies but allow the approach of smaller molecular substrates.

4.3. Comb-type polymer backbones

Comb-type polymers possess interesting characteristics because they are more well-defined than the hyperbranched polysaccharides discussed above. In addition, comb density, the length of the combs, and the bulkiness of the combs can be adjusted to confer different properties to the polymer and consequently also the conjugate. While indeed partial stretching of the main-chain of comb-type polymers is generally observed, the chains generally adopt conformations that render the overall system more compact than that adopted by linear polymers.⁸⁸ In this section, we survey the effect of comb density and comb length on conjugate properties given that, in all examples available, the chemical nature of the comb (MPEG) is the same.

In two separate studies, Ma and co-workers^89,90 have evaluated the effect of modification of ASNase with either linear P(VP-co-MA) or comb-shaped P(MPEG₅₀₀₀-g-VP-co-MA) (MA: maleic anhydride), that is, with or without polymeric side-chains. The authors observed that the degree of protein modification required for elimination of the antigenicity of ASNase decreased as the proportion of MPEG side-chains on the comb-type polymer increased. Complete elimination of antigenicity of ASNase was achieved by modification of 63 amino groups with linear P(VP-co-MA) in comparison to 30 amino groups for the comb-shaped P(MPEG₅₀₀₀-g-VP-co-MA). For comparison, complete elimination of antigenicity of ASNase modified with either linear or double branched MPEG₂ (2 × 5 kDa) is achieved only at higher degrees of enzyme modification and only low enzymatic activity is retained.^33,91 Interestingly, the comb-shaped polymer had a lower optimal activity temperature in comparison to the native enzyme, which may indicate that multi-point attachment between the protein and polymer is disfavored due to the bulky MPEG combs. Additionally, enzymatic activity remained higher when the polymer possessed a higher proportion of MPEG side-chains. Unfortunately, as the molecular weight of the conjugates have not been characterized in this study, it is not possible to further explain these results as it is unclear whether the number of polymer chains per ASNase molecule is altered as a function of increasing comb density.

Inada and co-workers^74,92 have modified both E. coli ASNase and BSA with both PM₁₃ and PM₁₀₀, which are comb-type polymers with 1.5–2 kDa MPEG side-chains and with overall molecular weights of 13 and 100 kDa, respectively (Fig. 3a). For ASNase, the authors observed that polymer molecular weight was an influential factor on antigenicity. That is, at a degree of modification at which no antigenicity of the conjugate was observed 46 and 30 lysine residues were modified by PM₁₃ and PM₁₀₀, respectively. This trend was also consistent for BSA for which antigenicity was lost following modification of 18 or 12 lysine residues, respectively (Fig. 3b). For both proteins, enzymatic activity was higher for PM₁₀₀ than for PM₁₃ or MPEG₂ (Table 1). Interestingly, and in contrast to the results of Ma and co-workers,^89,90 the conjugates had greater stability towards heat and denaturing conditions, and this as a function of polymer molecular weight. This result may perhaps stem from the shorter combs (i.e., 1.5–2 kDa) in PM₁₃ and PM₁₀₀ compared to the 5 kDa combs on the polymers of Ma and co-workers, which may have led to greater multi-point attachment. One could equally argue that the difference in length of the two combs is much less pronounced than the difference in the molecular weight of the two polymers, which could also be the dominant factor here.^89,90

Other comb-type conjugates with much smaller side-chains have been prepared by Lele et al.⁵³ by direct polymerization of PEGMA from a αCT macro-initiator. Increase of the molecular weight of the PPEGMA chain from ∼19 kDa to 160 kDa had only a small effect on activity, which indicates that individual comb-type polymer chains impose only a small diffusion barrier to small molecules. This may be an indication that the polymer is adopting a specific conformation that does not allow it to spread over the surface of the protein and interfere with its active site. In another example, Grover et al.⁶⁵ have modified bovine serum albumin with a single 7.1 kDa chain of PPEGA and have observed only a slight decrease in activity. This result should, however, be considered in the light of our previous comments on the enzymatic activity of BSA. Unfortunately, in both preceding examples, thermal stability and antigenicity have not been evaluated for comparison with the other conjugates from this section.

4.4. General trends

Based on the available literature it is not clear in what manner polymer backbone architecture can be exploited to enhance thermal stability and suppress antigenicity/proteolysis, without compromising enzymatic activity. It appears as though polymers with rigid backbones resulting either from branching, folding, or the presence of bulky side-chains influence to a lesser extent enzymatic activity of the conjugate than flexible linear polymers. This may be due to the lower ability of these polymers to spread over the surface of the enzyme and interfere with active regions. Also, the rigidity of the polymer may confer enhanced resistance to denaturation by heat as the polymer is likely to convey this property to the enzyme, thus stabilizing its structure and reducing its potential for denaturation, especially in the case of concurrent multi-point attachment.

5. Polymer side-chain functionality

In contrast to PEG, which is generally considered to be chemically inert under pharmaceutically relevant conditions, most of the natural and synthetic polymers in Tables 1–6 possess side-chains that are not inert and can thus interact either amongst themselves, with the solvent, with the protein, or with the enzymatic substrate. That is, non-covalent interactions such as hydrogen bonding (e.g., with polysaccharides, PHPMA, poly(N-vinylpyrrolidone) (PVP), etc.), electrostatic interactions (e.g., between positively charged lysine residues and ring-opened maleic anhydride repeat units), host–guest interactions (e.g., between the polymer and the protein, substrate, etc.), or other uncharacterized interactions also contribute to the overall properties of the enzyme. In this section, we successively discuss electrostatic interactions, host–guest interactions, and other less well-characterized interactions between a polymer and the enzyme or its enzymatic substrate and highlight the influence of these interactions on the bioactivity of the conjugate. We have found that it was not possible to distinguish properties related to hydrogen bonding from those stemming from covalent coupling and therefore have treated these together in Section 3.4. The specific effects discussed herein can be distinguished from polymer conjugation per se. Also, in certain cases, the authors have performed specific control experiments to identify the specificity of the phenomenon. Of worthy mention, in this section it is not unambiguous to distinguish whether the effect on bioactivity results from interaction between the polymer and the enzyme, between the polymer and the substrate, or to another uncharacterized phenomenon. In addition, the observed effects in subsection 5.3. tend to be relatively specific to a given polymer/enzyme/substrate system. Nevertheless, as ultimately the number of different polymers and enzymes typically used to prepare protein–polymer conjugates remains small, specific interactions such as those to be discussed in this section remain important and merit mention.

5.1. Electrostatic interactions

In Section 3.4 we have shown that neutral polymers conjugated to enzymes via multi-point attachment enhanced the thermal stability of the conjugate without altering its pH-activity profiles (Table 7). However, we have observed that in many instances where the polymer possessed a charge, pH-activity profiles of the conjugate were indeed altered (Table 7), indicating that in addition to the effects concurrent with polymer modification previously discussed, ionic interactions, whether between the polymer/enzyme or polymer/substrate, can also play a role in the characteristics of the conjugate. For example, Zhang et al.⁹³ have modified ASNase with sericin, a water-soluble polypeptide with molecular masses ranging from 20 to 60 kDa and composed of 15 types of amino acids, among which polar amino acids with hydroxyl, carboxyl, and amino groups such as serine, aspartic acid, and lysine account for 72% of all residues. The optimal operating pH of the resulting conjugate shifts considerably to 5 in comparison with pH 6–8 of the native form of the enzyme. In addition, the thermostability and resistance to trypsin digestion of the modified enzyme were greatly enhanced as compared with ASNase alone owing to the multi-point covalent coupling approach used to prepare the conjugate. Ma and co-workers^89,90 have modified ASNase with copolymers of PVP and maleic anhydride, both with and without pendant MPEG chains (i.e., comb-shaped architecture, discussed in Section 4.3). The pH optima of enzymatic activity of the modified ASNase was slightly more alkaline than the native enzyme (shifted by 0.4 and 0.8 units higher for copolymer with and without MPEG, respectively) which the authors attribute to the anionic microenvironment surrounding the enzyme. Qian et al.⁹⁴ have modified E. coli ASNase with N,O-carboxymethyl chitosan and have also noted that the modified enzyme showed slightly more alkaline pH optima than did the native enzyme. This may result from the charged nature of the polymer.

Lele et al.⁵³ have prepared conjugates of αCT by polymerization of sodium 4-styrenesulfonate (NaPS) and N,N-dimethylaminoethyl methacrylate (DMAEMA) from the protein macro-initiator. Interestingly, and in contrast to the preceding results for ASNase, comparable enzymatic activities are observed despite the significantly different structure and properties of these two polymers, indicative that for these particular systems the net charge of the conjugate is not a determining factor for enzymatic activity (at the tested pH). Based on this limited dataset, it is not possible to comment on whether this is a general phenomenon for αCT or whether it is simply valid for the above-mentioned conjugates.

Yan et al.⁹⁵ have coupled the thermosensitive zwitterionic polymer poly(3-dimethyl(methacryloyloxyethyl) ammonium propane sulfonate) (PDMAPS) to trypsin. The PDMAPS and the conjugate have their upper critical solution temperature at 14 and 16 °C, respectively. Investigation of the fluorescence of the tryptophan residues on the protein suggested that the conjugation did not lead to any significant change in the microenvironment of the tryptophan residues of the trypsin. In other words, the conjugated trypsin remained in its native tertiary conformation. Circular dichroism spectroscopy indicated that α-helical and β-sheet content of the conjugate were both slightly reduced. The optimal pH for activity was unchanged after conjugation, which the authors attribute to the fact that at pH 7 PDMAPS is neutral due to charge pairing. The conjugate had a smaller K_M as compared to that of the native trypsin, possibly due, as suggested by the authors, to enhanced hydrophobicity for the conjugated trypsin. Interestingly, the optimal temperature for proteolysis of casein of the conjugates increased from 20 °C (native) to 60 °C (conjugate), concurrent with a large increase of activity (well above the 100% activity of the native enzyme), though the reason for this is unclear.

In another example of conjugates prepared with zwitterionic polymers, Ishihara and co-workers^54,96 have modified papain with PMPC and have observed that the helix content of conjugated papain was slightly below that of native papain, indicating that conjugation slightly altered enzyme structure as a function of both the number of chains and polymer molecular weight. This change in enzyme conformation did not affect enzymatic activity as all conjugates, including a MPEG conjugate, had similar esterolytic activities (ca., 30–35%, Table 6). Unfortunately, the pH-dependent activity profile for these conjugates was not measured. Fujimura et al.⁹⁷ have covalently modified papain with different enteric coating polymers by means of carbodiimide coupling to the pendant carboxylic acid side-chains on the polymer. Heat stability of the resulting conjugates increased compared with that of native papain, and the conjugate had higher stability against water-miscible organic solvents and protein denaturing agents likely due to multi-point attachment. However, as the polymer itself is pH-sensitive, assessing the influence of the electrostatic interactions between the protein and the polymer on the optimum pH for activity is meaningless.

5.2. Host–guest interactions

Supramolecular interactions represent an interesting class of non-covalent interactions useful for stabilizing proteins. For instance, Fernandez et al.⁹⁸ have modified αCT with O-carboxymethyl-poly-β-cyclodextrin (Fig. 4) by coupling the pendant carboxylic acid groups to amino groups on the protein. This polymer has the potential to form stable inclusion complexes between cyclodextrin (βCD) units and hydrophobic compounds such as the substrate or certain amino acid residues located at the surface of the enzyme. The authors have found that the attached polymer served to maintain the active conformation of the enzyme after thermal treatment principally through covalent multi-point attachment of the polymer chains (via carbodiimide coupling), though multi-point salt bridges resulting from the polyanionic nature of the modifying polysaccharide, as well as through supramolecular interactions involving hydrophobic amino acid residues and the βCDs were speculated to contribute as well. This last phenomenon was verified by addition of 1-adamantanol, which forms a highly stable 1 : 1 complex with βCDs, resulting in a reduction of the protective effect of the polymer on the thermal resistance of the modified conjugate at 50 °C. Moreover, the half-life of the modified enzyme was only 1.5-fold increased in the presence of the guest 1-adamantanol with respect to the native counterpart, in contrast to an increase of about ∼12-fold without addition of the guest. The stabilization of enzymes through host–guest interactions by modification with βCDs has also been reported by others.^99–101 The thermal stabilization of trypsin using a βCD-modified dextran as additive (i.e., not covalently coupled to trypsin) has also been reported.¹⁰²

Fig. 4 (a) A schematic representation of O-carboxymethyl-poly-β-cyclodextrin prepared by polymerization of β-cyclodextrin (βCD) with epichlorohydrin.⁹⁸ (b) The monomeric form of bovine alpha-chymotrypsin (PDB entry 1YPH) with hydrophobic amino acids in blue (Val, Ile, and Leu dark; Ala, Phe, Cys, Met light), lysine residues in red, and the active catalytic site (His-57, Asp-102, and Ser-195) in magenta. The polymer in (a) was conjugated to the lysine residues on α-chymotrypsin. Host–guest interactions between the β-cyclodextrin units and hydrophobic amino acid residues on the surface of the enzyme resulted in enhanced stability of the conjugate towards heat. Part (a) is adapted from ref. 98. Protein image produced with PyMol v.1.1.

5.3. Other interactions or effects

In addition to the specific examples above, the functional groups along the backbone of the polymer may influence the polarity, diffusivity, etc. of the microenvironment surrounding the enzyme in a less precise manner, thus affecting how the substrate interacts with the latter. In this subsection we summarize examples in which the nature of the polymer has a distinct effect on enzymatic activity of the conjugate, whether this underlying phenomenon has been elucidated or not.

von Specht and Brendel¹⁰³ have covalently modified trypsin with PVP by multi-point attachment by prior opening of a certain percentage of the lactam side-chains. The resulting conjugate showed an increase in specific activity compared to the native enzyme towards low molecular weight substrates. However, activity towards large substrates such as azocasein was much reduced. This difference of activity indicates that the main changes which are caused by binding short chains of PVP are of steric origin. The authors attributed the increased activity of trypsin towards small substrates to an increased local concentration of the latter stemming from the known binding ability of PVP towards small molecule dyes and drugs. Ding et al.⁵² have prepared conjugates of trypsin and semi-telechelic PNIPAAm. The authors observed that the esterase activity of the conjugates increased in relation to that of the native protein. Furthermore an increase of activity with the extent of polymer conjugation was observed. These effects were also observed in the amidase activity of the conjugate. The authors speculated that this could be related to the change in microenvironment of the enzyme's active site, leading to enhanced partitioning of substrate in the polymer phase. Lee and Park¹⁰⁴ have prepared conjugates of trypsin with a semi-telechelic copolymer composed of NIPAAm and a glucose monomer and bearing a terminal carboxylic acid group, which was used for coupling to protein. By considering the weight proportion of enzyme within the conjugate, the amidase activity of the enzymes appeared to be enhanced compared to the native enzyme. Interestingly, the conjugated enzymes demonstrated very peculiar enzyme activity-temperature profiles, with two apparent optimal temperatures, indicating that both temperature-controlled collapse and flocculation of the copolymers around the enzyme surface modulated the mass transfer rates of substrate to the active site of the enzyme.

Interestingly, lysozyme modified with 35.6 kDa PMPC in a random fashion exhibited ∼100% activity towards the macromolecular substrate Micrococcus Luteus, which contrasts with the known deactivation of this enzyme upon random conjugation of a polymer to lysine residues (Table 4).⁶⁶ Whether this is a specific effect of the zwitterionic polymer or due to the choice of a different substrate (M. Lysodeikticus is a more generally used substrate in Table 4) cannot be established herein. Yuan et al.¹⁰⁵ have exploited the self-assembly characteristics of a lysozyme/MPEG–poly(aspartic acid)) (MPEG₅₀₀₀-PASP₃₄) mixture to form polyion complex (PIC) micelles, which were subsequently cross-linked by addition of gluteraldehyde. This step promoted reaction between the amino groups of lysozyme and the ω-amino end-group of the polymer within the core of the micelle. The authors have demonstrated the effect of substrate sequestering on the enzymatic activity towards a synthetic small-molecule substrate able to penetrate into the core of the micelle. In comparison to the native enzyme, the conjugates displayed significantly enhanced activity, independently of the amount of gluteraldehyde added for cross-linking indicating that, within the experiments performed, core-cross-linking did not inhibit the enzymatic activity of the entrapped lysozyme. Danial et al.¹⁰⁶ have prepared complex coacervate core micelles with a lysozyme-modified corona by coupling the latter to the end of a poly(2-vinyl pyridine)-b-PEG block copolymer. The enzymatic activity of the lysozyme-micelle conjugates towards M. lysodeikticus was comparable to that of free lysozyme, though decreased when the number of enzyme molecules per micelle increased. This was attributed to sequestration of a certain number of enzymes within the core of the micelle, where interaction with the substrate was hindered.

Miyamoto et al.¹⁰⁷ have increased the hydrophobicity of PMPC by copolymerization with n-butyl methacrylate (BMA) in differing proportions and have conjugated the resulting polymers to papain (Fig. 5). The authors observed a loss of α-helical content of the papain only for the most hydrophobic block copolymer prepared from these two monomers. Concurrently, enzymatic activity decreased with increasing amounts of BMA in the copolymer, which might be due to enhanced restriction of substrate diffusion to the enzyme. These polymers nevertheless stabilized the protein due to prevention of auto-digestion.

Fig. 5 (a) Copolymers of the zwitterionic monomer MPC with the hydrophobic monomer BMA were conjugated to lysine residues (red) on papain (b, PDB entry 1PPP). The active site of the enzyme (Cys-25, His-159, and Asp-158) is shown in magenta. The activity of the enzyme decrease with increased BMA content.¹⁰⁷ Protein image produced with PyMol v.1.1.

5.4. General trends

Within the examples examined, it appears as though polymers bearing charged functional side-chains affect the pH-dependent activity profiles of enzymes such as ASNase and trypsin. This was not the case, however, for αCT whose pH-dependent activity was not affected by conjugation of charged polymers. Interestingly, the zwitterionic polymer PDMAPS did not alter the pH optima of activity of trypsin. Host–guest interactions between polymeric βCDs and hydrophobic residues on αCT or trypsin appear to effectively improve thermal stability, whether the polymer is covalently coupled to the enzyme or not. This phenomenon is readily distinguished from all other interactions by addition of 1-adamantanol. The last subsection has highlighted that the functional side-chains of the polymer can contribute to phenomena which have a very significant effect on bioactivity. This effect can in fact lead to a pronounced improvement of catalytic activity in comparison to the native enzyme. Overall, this section demonstrates that polymer side-chains functionality must be considered either in conjugate design (for the limited trends above) or when interpreting conjugate properties.

6. Polymer biodegradability

Biodegradability is an important feature of protein–polymer conjugates designed for use in humans. Increasing the molecular weight of therapeutic proteins through polymer conjugation is an easy way of avoiding clearance by the kidneys, though accumulation of large, non-biodegradable components in the liver may lead to undesirable side-effects. Understanding the mechanism through which the protein–polymer conjugate will decompose over time is important for understanding how its activity will evolve over this period, and for predicting its ultimate fate in the body. Biodegradability can be achieved in a number of ways.¹⁰⁸ In the simplest case, the polymer can be biodegradable along its main-chain, leading to a progressive reduction in molecular weight and production of small molecules. Polypeptides and polysaccharides are examples of such polymers that may degrade under appropriate hydrolytic and/or enzymatic conditions. Alternatively, a non-biodegradable polymer may be conjugated to the enzyme via a labile linker, which, upon biodegradation, releases the intact polymer chains. In this case, care must be taken that the individual polymer chains have molecular weights low enough to be eliminated by the kidneys. Conjugates containing MPEG coupled to the protein via a cleavable linker are examples of such systems. To a certain extent, all polymers linked to the protein via a potentially labile bond, such as an ester or an amide bond, also fall into this category. Lastly, the polymer may hydrolyze along its side-chain, thus progressively reducing the molecular weight. Polymers with large side-chains such as ca. PM₁₃ or PM₁₀₀ fall into this category as release of high molecular MPEG upon hydrolysis of the ester bond which binds it to the main-chain would dramatically reduce the weight of the parent polymer. In this section, we discuss the effect of these three forms of biodegradation on enzymatic activity.

6.1. Main-chain biodegradable polymers

Polypeptides are susceptible to proteolysis and therefore fall into the category of main-chain biodegradable polymers. Within the literature examined herein, no specific mention is made of attempting to selectively remove polypeptides covalently coupled to enzymes. However, in certain cases, the stability of the conjugates in the presence of proteolytic enzymes is reported and the enzymatic activity reported as a function of time. This experiment indirectly examines the ability to selectively degrade the conjugated polypeptide chains versus the enzyme within its core. For instance, Uren and Ragin⁴⁸ have prepared conjugates of poly(DL-alanine) with ASNases and have evaluated different proteolytic conditions for assaying conjugate stability. The authors observed that, over a one hour period, the conjugate was totally resistant to trypsin, while 50% of enzyme activity was lost for the native protein. Therefore, under these conditions, any possible proteolysis of poly(DL-alanine) had little or no effect on the activity of the conjugate. In contrast, upon incubation of the conjugate with αCT, enzymatic activity was reduced, though to a significantly lesser extent that for native ASNase. Given that, a priori, poly(DL-alanine) does not contain aromatic residues which are the typical substrate for αCT, it is possible that the primary effect observed is the retarded degradation of ASNase due to the imposed steric barrier of the polymer. These results indicate that it is not possible to selectively degrade the poly(DL-alanine) with αCT without compromising ASNase activity. In another example, Poznansky et al.⁸⁷ have evaluated the proteolytic stability of ASNase modified with albumin, a polypeptide which is susceptible to tryptic digestion. Interestingly, over the first half hour after exposure to trypsin, the activity of the enzyme increased (very slightly), which may be an indication that biodegradation of the protective shell around the enzyme is occurring and thereby increasing the activity of the enzyme. This increase, however, is minor and is followed by a more rapid decrease of activity over the next four hours, indicative of ASNase degradation. Shen and co-workers^72,93 have evaluated the stability of ASNase modified with silk fibroin and sericin and have also concurrently observed only a decrease of enzymatic activity over time, indicative that the conjugated proteins cannot be selectively degraded without compromising ASNase activity.

In contrast to polypeptides, polysaccharides may be degraded under conditions which, in principle, should not affect the enzyme. Marshall and Rabinowitz⁸¹ have evaluated the biodegradation of a trypsin–dextran conjugate in the presence of dextranase. The authors interestingly observed no change in esterolytic or caseinolytic activity following treatment with the enzyme. In addition, the authors observed that the susceptibility of the conjugates to inhibition with ovomucoid increased following dextranase treatment, indicating that removal of dextran increased the accessibility of the inhibitor to the active site of the conjugated enzyme. Consequently, the authors concluded that the non-recovery of activity of the conjugate following dextranase treatment is due to carbohydrate attachment per se rather than concomitant burying of enzyme's active site by polymer chains. More recently, Duncan et al.⁷⁹ have evaluated the biodegradability of trypsin–dextrin conjugates by α-amylase. The enzymatic activity of the conjugate increased upon exposure to the enzyme, though the extent of regeneration was difficult to interpret. The authors indicated that this would be related to the rate of polymer degradation, the liberation of active protein with little or no glucose oligomers attached, and maintenance of protein integrity during conjugation and subsequent biodegradation.

6.2. Labile linkers for non-biodegradable polymers

A relatively recent approach for regenerating enzymatic activity of protein–polymer conjugates in the body is to introduce a cleavable linker between the polymer and the enzyme.¹⁰⁹ Examples of conjugates of lysozyme with MPEG containing a labile linker are found in Table 4. Lysozyme is an excellent model protein to evaluate this phenomenon because conjugation of a single 5 kDa MPEG chain to a lysine residue on this protein has been shown to completely eliminate enzymatic activity.^{60,61,110,111} These linkers are intercalated between the polymer and the protein and are designed to release these two moieties by hydrolysis in the body. This approach is, in principle, amenable to any non-biodegradable polymer. These authors have exploited cleavable linkers to reactivate the enzyme, which in most cases reached ∼100% of the activity of the native enzyme after an appropriate time of incubation. One exception is the conjugate prepared by Roberts and Harris,¹¹⁰ which only achieved 60% reactivation. This may be due to the fact that in this case linker cleavage is not “traceless” in as much as the lysine residue is not recovered but remained modified (with a small molecule) following linker cleavage. This has been shown in a study by Zhao et al.⁶¹ who have modified the amino groups on lysozyme with small molecules such as maleic anhydride, resulting in a reduction of activity of 60% and thereby demonstrating that the attachment of a small molecule to an amine of lysozyme can also inhibit activity. The authors further speculated that possible denaturation and conformational changes in heavily PEGylated molecules, which are not reversible upon release of the polymer contribute to this phenomenon.

6.3. Side-chain biodegradable polymers

Vinyl polymers are generally considered to be non-biodegradable along their main-chain. One exception, though not treated herein, are poly(alkyl cyanoacrylate)s. However, the functional side-chains of main-chain non-biodegradable polymers are generally attached to the backbone by ester or amide bonds, which have the possibility of degrading inside the body. On the one hand, this process is undesirable because of complications arising from the need to evaluate the potential bioactivity of the biodegradation products. On the other, this process can, over time, dramatically reduce the molecular weight of the polymer, especially when the side-groups have large molecular weights. This holds particularly true for comb-type polymers which possess large side-chains such as those employed by Qian et al.⁸⁹ and Lele et al.⁵³ (5 and 1.1 kDa, respectively). Clearly, the amide and ester groups present on typical (meth)acrylate or (meth)acrylamide polymers are expected to be quite stable during the life span of the conjugate and thus should not affect the activity of the conjugate over this period. In terms of ultimate fate however, there is unfortunately very little information available on the rates of non-specific hydrolysis of polymer side-chains in the body (or in simulated conditions) or on the biological fate in vivo of many common polymers, whether biodegradable or not.^112–119 Further studies will become necessary as future research in this field is pursued.

6.4. General trends

In general, in the examples cited, polypeptide biodegradation resulted in little or no recovery of enzyme activity, which may result from the similarity between polypeptide and enzyme degradation conditions. The selective biodegradation of polysaccharides, however, is possible and lead in certain cases to increased accessibility of small-molecules to the active site of the enzyme, though this did not always correlate with a complete recovery of activity. This phenomenon was also observed for when certain “non-traceless” cleavable linkers were used to prepare biodegradable MPEG–lysozyme conjugates.

7. Outlook

Through this review article we've highlighted trends observed using new protein–polymer conjugation strategies, with particular emphasis on how they relate to enzymatic activity. It should become apparent that modern tools for preparing protein–polymer conjugates allow for exceptional freedom in conjugate design and as such it is somewhat difficult to find comparable conjugates within the literature. In addition, there are very few systematic studies which have been performed to assess the individual contributions of many of the factors discussed herein, which makes it very difficult to make general statements. Owing to the specificity and dissimilarity of trends observed and summarized in Table 7, we feel it would be premature to provide general guidelines regarding optimum molecular weight, residue for modification, distance from active sites, etc., for maintaining catalytic activity of conjugates. We have however provided a summary of the specific effect of state-of-the-art protein modifications on activity and have summarized trends observed for each protein in Table 7.

As more and more conjugates are prepared in the coming years, the generality of the statements made herein, for select proteins, may be verified or negated. One clear point which is to be made with this review is that no one factor is uniquely responsible for conjugate properties, rather the cumulative effect of the applicable factors described herein governs its ultimate properties. It is our hope with this contribution to summarize the state-of-the-art in this rapidly expanding field with the desire to provide (at least tentative) guidelines for preparing future functional protein–polymer constructs.

One final topic which we have not touched upon in this contribution is that of immunogenicity. Immunogenicity is an important and intrinsic property of the polymer and/or the conjugate and its evaluation must be made concurrently with the development of such systems, independently of the polymer used. Even PEG, with the plethora of data to support its use for therapeutic purposes, cannot escape this requirement especially as recent studies are beginning to identify anti-PEG antibodies in animals.^120–122

8. List of abbreviations for tables

AADG	acrylamido-2-deoxy-D-glucose
ABr	via acid bromide
Alk	alkylation with α-bromoester
AzoT	via azo trypsin
bisAlk	bis-alkylation via a three-carbon bridge
BMA	n-butyl methacrylate
CAP	cellulose acetate phthalate
CDI	via carbodiimide
CMC	carboxymethyl cellulose
CM-PβCD	O-carboxymethyl-poly-β-cyclodextrin
CNBr	cyanogen bromide
Eudragit-S	1 : 2 methacrylic acid-co-methyl methacrylate
HP-55	hydroxy propylmethylcellulose phthalate
Hyd	via hydrazide
GA	via gluteraldehyde
GEMA	glucosyoxylethyl methacrylate
MA	maleic anhydride
MAA	methacrylic acid
Mal	via Michael addition with maleimide
MMA	methyl methacrylate
MPEG	α-methoxy-poly(ethylene glycol)
MPEG-deg	MPEG with degradable linker
MPEG2	double branched MPEG
MPM-06	methylacrylate-methacrylic acid-methylmethacrylate copolymer
n.c.	not characterized
NC	non-covalent
NCA	via α-amino acid N-carboxy anhydride polymerization
n.g.	not given
NHS	via N-hydroxysuccinimide active ester
NIPAAm	N-isopropylacrylamide
N,O-CM-Chit	N,O-CM-chitosan
NPC	p-nitrophenylcarbonate
OTs	via tosylate
PAGM	poly(aminoglucose methacrylate)
PAsp	poly(aspartic acid)
PCVBPC	poly(3-carbamoyl-1-(p-vinylbenzyl)pyridinium chloride)
PDMAEMA	poly(N,N-dimethylaminoethyl methacrylate)
PDMAPS	poly(3-dimethyl(methacryloyloxyethyl) ammonium propane sulfonate)
PEG	poly(ethylene glycol)
PEO	poly(ethylene oxide)
PHPMA	poly(2-hydroxypropyl methacrylamide)
PM13	copolymer of poly(oxyethylene) allyl methyl diether and maleic anhydride
PM100	copolymer of poly(oxyethylene) 2-methyl-2-propenyl methyl diether and maleic anhydride
PMPC	poly(2-methacryloyloxyethyl phosphorylcholine)
PNaPS	poly(sodium 4-styrenesulfonate)
PNIPAAm	poly(N-isopropylacrylamide)
pNP	via p-nitrophenyl ester
POZ	poly(2-ethyl-oxazoline)
PPEGA	poly(polyethylene glycol monomethyl ether acrylate)
PPEGMA and PPEGMA475	poly(polyethylene glycol monomethyl ether methacrylate)
PSucr	polymeric sucrose
PVP	poly(vinyl pyrrolidone)
RA	via reductive alkylation
RPE and REP	block copolymers of ethylene oxide and propylene oxide, (R: radical, P: propylene oxide, E: ethylene oxide)
SC	via succinimidyl carbonate
SS	via disulfide exchange
TCT	via 2,4,6-trichloro[1,3,5]-triazine
TGase	transglutaminase
TT	via thiazolidine-2-thione coupling
VP	vinyl pyrrolidone
VS	via vinyl sulfone.

Acknowledgements

H.-A.K. thanks the Swiss National Science Foundation, the NCCR Nanoscale Science, and the European Commission for supporting his laboratory's activities on peptide/protein–synthetic polymer conjugates. M.A.G. gratefully acknowledges a scholarship from the Fonds Québecois de la Recherche sur la Nature et les Technologies (Quebec, Canada) for funding his postdoctoral research on protein–polymer conjugates with H.-A. K from 2007–2009.

Notes and references

1. E. Pedone and S. Brocchini, React. Funct. Polym., 2006, 66, 167–176.
2. G. Pasut and F. M. Veronese, Prog. Polym. Sci., 2007, 32, 933–961.
3. F. M. Veronese, Biomaterials, 2001, 22, 405–417.
4. G. Pasut, M. Sergi and F. M. Veronese, Adv. Drug Delivery Rev., 2008, 60, 69–78.
5. J. M. Harris, N. E. Martin and M. Modi, Clin. Pharmacokinet., 2001, 40, 539–551.
6. G. Pasut and F. M. Veronese, Adv. Drug Delivery Rev., 2009, 61, 1177–1188.
7. S. Brocchini, A. Godwin, S. Balan, J.-w. Choi, M. Zloh and S. Shaunak, Adv. Drug Delivery Rev., 2008, 60, 3–12.
8. A. J. de Graaf, M. Kooijman, W. E. Hennink and E. Mastrobattista, Bioconjugate Chem., 2009, 20, 1281–1295.
9. H. B. F. Dixon, J. Protein Chem., 1984, 3, 99–108.
10. M. A. Gauthier and H.-A. Klok, Chem. Commun., 2008, 2591–2611.
11. P. Thordarson, B. Le Droumaguet and K. Velonia, Appl. Microbiol. Biotechnol., 2006, 73, 243–254.
12. H.-A. Klok, J. Polym. Sci., Part A: Polym. Chem., 2005, 43, 1–17.
13. J. M. Antos and M. B. Francis, Curr. Opin. Chem. Biol., 2006, 10, 253–262.
14. D. R. W. Hodgson and J. M. Sanderson, Chem. Soc. Rev., 2004, 33, 422–430.
15. R. E. Connor and D. A. Tirrell, J. Macromol. Sci., Polym. Rev., 2007, 47, 9–28.
16. A. J. Link, M. L. Mock and D. A. Tirrell, Curr. Opin. Biotechnol., 2003, 14, 603–609.
17. J. C. M. van Hest, J. Macromol. Sci., Polym. Rev., 2007, 47, 63–92.
18. K. L. Heredia and H. D. Maynard, Org. Biomol. Chem., 2007, 5, 45–53.
19. H. G. Börner and H. Schlaad, Soft Matter, 2007, 3, 394–408.
20. D. W. P. M. Löwik, L. Ayres, J. M. Smeenk and J. C. M. van Hest, Adv. Polym. Sci., 2006, 202, 19–52.
21. L. A. Canalle, D. W. P. M. Löwik and J. C. M.v. Hest, Chem. Soc. Rev., 2010, 39, 329–353.
22. H.-A. Klok, Macromolecules, 2009, 42, 7990–8000.
23. J. Nicolas, G. Mantovani and D. M. Haddleton, Macromol. Rapid Commun., 2007, 28, 1083–1111.
24. V. Coessens, T. Pintauer and K. Matyjaszewski, Prog. Polym. Sci., 2001, 26, 337–377.
25. G. Moad, Y. K. Chong, A. Postma, E. Rizzardo and S. H. Thang, Polymer, 2005, 46, 8458–8468.
26. A. Hirao and M. Hayashi, Acta Polym., 1999, 50, 219–231.
27. B. Le Droumaguet and J. Nicolas, Polym. Chem., 2010, 1 10.1039/b9py00363k.
28. Y. Inada, M. Furukawa, H. Sasaki, Y. Kodera, M. Hiroto, H. Nishimura and A. Matsushima, Trends Biotechnol., 1995, 13, 86–91.
29. V. Gaberc-Porekar, I. Zore, B. Podobnik and V. Menart, Curr. Opin. Drug Discovery Dev., 2008, 11, 242–250.
30. A. Fontana, B. Spolaore, A. Mero and F. M. Veronese, Adv. Drug Delivery Rev., 2008, 60, 13–28.
31. A. S. Hoffman and P. S. Stayton, Prog. Polym. Sci., 2007, 32, 922–932.
32. R. Eisenthal, M. J. Danson and D. W. Hough, Trends Biotechnol., 2007, 25, 247–249.
33. Y. Ashihara, T. Kono, S. Yamazaki and Y. Inada, Biochem. Biophys. Res. Commun., 1978, 83, 385–391.
34. A. Matsushima, H. Nishimura, Y. Ashihara and Y. Inada, Chem. Lett., 1980, 773–776.
35. J.-F. Zhang, L.-Y. Shi and D.-Z. Wei, Biotechnol. Lett., 2004, 26, 753–756.
36. A. L. Soares, G. M. Guimarães, B. Polakiewicz, R. N. d. M. Pitombo and J. Abrahão-Neto, Int. J. Pharm., 2002, 237, 163–170.
37. C. Monfardini, O. Schiavon, P. Caliceti, M. Morpurgo, J. M. Harris and F. M. Veronese, Bioconjugate Chem., 1995, 6, 62–69.
38. F. M. Veronese, C. Monfardini, P. Caliceti, O. Schiavon, M. D. Scrawen and D. Beer, J. Controlled Release, 1996, 40, 199–209.
39. N. Verma, K. Kumar, G. Kaur and S. Anand, Crit. Rev. Biotechnol., 2007, 27, 45–62.
40. J. A. Rodríguez-Martínez, I. Rivera-Rivera, R. J. Solá and K. Griebenow, Biotechnol. Lett., 2009, 31, 883–887.
41. I. J. Castellanos, W. Al-Azzam and K. Griebenow, J. Pharm. Sci., 2005, 94, 327–340.
42. H.-C. Chiu, S. Zalipsky, P. Kopečková and J. Kopeček, Bioconjugate Chem., 1993, 4, 290–295.
43. H. F. Gaertner and A. J. Puigserver, Enzyme Microb. Technol., 1992, 14, 150–155.
44. L. Goldstein, Biochemistry, 1972, 11, 4072–4084.
45. A. Abuchowski and F. F. Davis, Biochim. Biophys. Acta, Protein Struct., 1979, 578, 41–46.
46. B. Treetharnmathurot, C. Ovartlarnporn, J. Wungsintaweekul, R. Duncan and R. Wiwattanapatapee, Int. J. Pharm., 2008, 357, 252–259.
47. T. So, T. Ueda, Y. Abe, T. Nakamata and T. Imoto, J. Biochem., 1996, 119, 1086–1093.
48. J. R. Uren and R. C. Ragin, Cancer Res., 1979, 39, 1927–1933.
49. T. Yoshimura, A. Imanishi and T. Isemura, J. Biochem., 1968, 63, 730–737.
50. Z.-R. Lu, P. Kopečková, Z. Wu and J. Kopeček, Bioconjugate Chem., 1998, 9, 793–804.
51. D. Oupický, K. Ulbrich and B. Říhová, J. Bioact. Compat. Polym., 1999, 14, 213–231.
52. Z. Ding, G. Chen and A. S. Hoffman, J. Biomed. Mater. Res., 1998, 39, 498–505.
53. B. S. Lele, H. Murata, K. Matyjaszewski and A. J. Russell, Biomacromolecules, 2005, 6, 3380–3387.
54. D. Miyamoto, J. Watanabe and K. Ishihara, Biomaterials, 2004, 25, 71–76.
55. R. J. Solá and K. Griebenow, FEBS J., 2006, 273, 5303–5319.
56. S. D. Varfolomeev and K. G. Gurevich, Russ. Chem. Bull., 2001, 50, 1709–1717.
57. G. J. Palm, J. Lubkowski, C. Derst, S. Schleper, K.-H. Röhm and A. Wlodawer, FEBS Lett., 1996, 390, 211–216.
58. S. Balan, J.-w. Choi, A. Godwin, I. Teo, C. M. Laborde, S. Heidelberger, M. Zloh, S. Shaunak and S. Brocchini, Bioconjugate Chem., 2007, 18, 61–76.
59. T. Higashi, F. Hirayama, S. Yamashita, S. Misumi, H. Arima and K. Uekama, Int. J. Pharm., 2009, 374, 26–32.
60. R. B. Greenwald, K. Yang, H. Zhao, C. D. Conover, S. Lee and D. Filpula, Bioconjugate Chem., 2003, 14, 395–403.
61. H. Zhao, K. Yang, A. Martinez, A. Basu, R. Chintala, H.-C. Liu, A. Janjua, M. Wang and D. Filpula, Bioconjugate Chem., 2006, 17, 341–351.
62. K. L. Heredia, D. Bontempo, T. Ly, J. T. Byers, S. Halstenberg and H. D. Maynard, J. Am. Chem. Soc., 2005, 127, 16955–16960.
63. C. Boyer, V. Bulmus, J. Liu, T. P. Davis, M. H. Stenzel and C. Barner-Kowollik, J. Am. Chem. Soc., 2007, 129, 7145–7154.
64. P. De, M. Li, S. R. Gondi and B. S. Sumerlin, J. Am. Chem. Soc., 2008, 130, 11288–11289.
65. G. N. Grover, S. N. S. Alconcel, N. M. Matsumoto and H. D. Maynard, Macromolecules, 2009, 42, 7657–7663.
66. J.-H. Seo, R. Matsuno, Y. Lee, M. Takai and K. Ishihara, Biomaterials, 2009, 30, 4859–4867.
67. J. Córdova, J. D. Ryan, B. B. Boonyaratanakornkit and D. S. Clark, Enzyme Microb. Technol., 2008, 42, 278–283.
68. Y. Ito, M. Kotoura, D.-J. Chung and Y. Imanishi, Bioconjugate Chem., 1993, 4, 358–361.
69. J. Liu, V. Bulmus, D. L. Herlambang, C. Barner-Kowollik, M. H. Stenzel and T. P. Davis, Angew. Chem., Int. Ed., 2007, 46, 3099–3103.
70. V. V. Mozhaev, N. S. Melik-Nubarov, M. V. Sergeeva, V. Šikšnis and K. Martinek, Biocatal. Biotransform., 1990, 3, 179–187.
71. T. Yoshimoto, K. Takahashi, A. Ajima, A. Matsushima, Y. Saito, Y. Tamaura and Y. Inada, FEBS Lett., 1985, 183, 170–172.
72. Y.-Q. Zhang, W.-L. Zhou, W.-D. Shen, Y.-H. Chen, X.-M. Zha, K. Shirai and K. Kiguchi, J. Biotechnol., 2005, 120, 315–326.
73. I. Vīna, A. Karsakevich and M. Bekers, J. Mol. Catal. B: Enzym., 2001, 11, 551–558.
74. Y. Kodera, T. Sekine, T. Yasukohchi, Y. Kiriu, M. Hiroto, A. Matsushima and Y. Inada, Bioconjugate Chem., 1994, 5, 283–286.
75. H. K. Kim and T. G. Park, Enzyme Microb. Technol., 1999, 25, 31–37.
76. P. Wang, T. G. Hill, C. A. Wartchow, M. E. Huston, L. M. Oehler, M. B. Smith, M. D. Bednarski and M. R. Callstrom, J. Am. Chem. Soc., 1992, 114, 378–380.
77. A. Lääne, V. Chytrý, M. Haga, P. Sikk, A. Aaviksaar and J. Kopeček, Collect. Czech. Chem. Commun., 1981, 46, 1466–1473.
78. P. V. Sundaram and R. Venkatesh, Protein Eng., Des. Sel., 1998, 11, 699–705.
79. R. Duncan, H. R. P. Gilbert, R. J. Carbajo and M. J. Vicent, Biomacromolecules, 2008, 9, 1146–1154.
80. B. Treetharnmathurot, L. Dieudonné, E. L. Ferguson, D. Schmaljohann, R. Duncan and R. Wiwattanapatapee, Int. J. Pharm., 2009, 373, 68–76.
81. J. J. Marshall and M. L. Rabinowitz, J. Biol. Chem., 1976, 251, 1081–1087.
82. M. L. Villalonga, R. Villalonga, L. Mariniello, L. Gómez, P. Di Pierro and R. Porta, World J. Microbiol. Biotechnol., 2006, 22, 595–602.
83. N. Rajalakshmi and P. V. Sundaram, Protein Eng., Des. Sel., 1995, 8, 1039–1047.
84. D. Venturoli and B. Rippe, Am. J. Physiol.: Renal Physiol., 2005, 288, F605–613.
85. S. S. Stivala, J. Kimura and L. Reich, Thermochim. Acta, 1981, 50, 111–122.
86. T. C. Le, B. D. Todd, P. J. Daivis and A. Uhlherr, J. Chem. Phys., 2009, 131, 044902–044910.
87. M. J. Poznansky, M. Shandling, M. A. Salkie, J. Elliott and E. Lau, Cancer Res., 1982, 42, 1020–1025.
88. Y.-J. Sheng, K.-L. Cheng and C.-C. Ho, J. Chem. Phys., 2004, 121, 1962–1968.
89. G. Qian, J. Ma, J. Zhou, B. He and D. Wang, Polym. Adv. Technol., 1997, 8, 581–586.
90. G. Qian, J. Ma, J. Zhou and B. He, React. Funct. Polym., 1997, 32, 117–121.
91. Y. Kamisaki, H. Wada, T. Yagura, A. Matsushima and Y. Inada, J. Pharmacol. Exp. Ther., 1981, 216, 410–414.
92. H. Sasaki, Y. Ohtake, A. Matsushima, M. Hiroto, Y. Kodera and Y. Inada, Biochem. Biophys. Res. Commun., 1993, 197, 287–291.
93. Y.-Q. Zhang, M.-L. Tao, W.-D. Shen, J.-P. Mao and Y.-h. Chen, J. Chem. Technol. Biotechnol., 2006, 81, 136–145.
94. G. Qian, J. Zhou, J. Ma, D. Wang and B. He, Artif. Cells, Blood Substitutes, Biotechnol., 1996, 24, 567–577.
95. M. Yan, J. Ge, W. Dong, Z. Liu and P. Ouyang, Biochem. Eng. J., 2006, 30, 48–54.
96. D. Miyamoto, J. Watanabe and K. Ishihara, J. Appl. Polym. Sci., 2004, 91, 827–832.
97. M. Fujimura, T. Mori and T. Tosa, Biotechnol. Bioeng., 1987, 29, 747–752.
98. M. Fernández, M. L. Villalonga, A. Fragoso, R. Cao, M. Baños and R. Villalonga, Process Biochem., 2004, 39, 535–539.
99. R. Villalonga, M. Fernández, A. Fragoso, R. Cao, L. Mariniello and R. Porta, Biotechnol. Appl. Biochem., 2003, 38, 53–59.
100. M. Fernández, M. L. Villalonga, A. Fragoso, R. Cao and R. Villalonga, Biotechnol. Appl. Biochem., 2002, 36, 235–239.
101. M. Fernández, A. Fragoso, R. Cao and R. Villalonga, Process Biochem., 2005, 40, 2091–2094.
102. M. Fernández, M. L. Villalonga, J. Caballero, A. Fragoso, R. Cao and R. Villalonga, Biotechnol. Bioeng., 2003, 83, 743–747.
103. B.-U. von Specht and W. Brendel, Biochim. Biophys. Acta, Enzymol., 1977, 484, 109–114.
104. H. Lee and T. G. Park, Biotechnol. Prog., 1998, 14, 508–516.
105. X. Yuan, Y. Yamasaki, A. Harada and K. Kataoka, Polymer, 2005, 46, 7749–7758.
106. M. Danial, H.-A. Klok, W. Norde and M. A. Cohen Stuart, Langmuir, 2007, 23, 8003–8009.
107. D. Miyamoto, J. Watanabe and K. Ishihara, J. Appl. Polym. Sci., 2005, 95, 615–622.
108. W. R. Gombotz and D. K. Pettit, Bioconjugate Chem., 1995, 6, 332–351.
109. D. Filpula and H. Zhao, Adv. Drug Delivery Rev., 2008, 60, 29–49.
110. M. J. Roberts and J. M. Harris, J. Pharm. Sci., 1998, 87, 1440–1445.
111. S. Lee, R. B. Greenwald, J. McGuire, K. Yang and C. Shi, Bioconjugate Chem., 2001, 12, 163–169.
112. N. Bertrand, J. G. Fleischer, K. M. Wasan and J.-C. Leroux, Biomaterials, 2009, 30, 2598–2605.
113. Y. Murakami, Y. Tabata and Y. Ikada, Drug Delivery, 1997, 4, 23–31.
114. Y. Takakura, T. Fujita, M. Hashida and H. Sezaki, Pharm. Res., 1990, 7, 339–346.
115. T. Yamaoka, Y. Tabata and Y. Ikada, J. Pharm. Pharmacol., 1995, 47, 479–486.
116. L. W. Seymour, R. Duncan, J. Strohalm and J. Kopeček, J. Biomed. Mater. Res., 1987, 21, 1341–1358.
117. L. W. Seymour, Y. Miyamoto, H. Maeda, M. Brereton, J. Strohalm, K. Ulbrich and R. Duncan, Eur. J. Cancer, 1995, 31, 766–770.
118. T. Yamaoka, Y. Tabata and Y. Ikada, J. Pharm. Sci., 1994, 83, 601–606.
119. T. Yamaoka, Y. Tabata and Y. Ikada, J. Pharm. Sci., 1995, 84, 349–354.
120. X. Y. Wang, T. Ishida and H. Kiwada, J. Controlled Release, 2007, 119, 236–244.
121. A. Judge, K. McClintock, J. R. Phelps and I. MacLachlan, Mol. Ther., 2006, 13, 328–337.
122. K. Środa, J. Rydlewski, M. Langner, A. Kozubek, M. Grzybek and A. F. Sikorski, Cell. Mol. Biol. Lett., 2005, 10, 37–47.

---