A universal fluorescent acceptor for high-performance liquid chromatography analysis of pro- and eukaryotic polysialyltransferases

Analytical Biochemistry 427 (2012) 107–115

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A universal fluorescent acceptor for high-performance liquid chromatography analysis of pro- and eukaryotic polysialyltransferases Timothy G. Keys 1, Friedrich Freiberger 1, Jörg Ehrit, Jonas Krueger, Katinka Eggers, Falk F.R. Buettner, Rita Gerardy-Schahn ⇑ Institute for Cellular Chemistry, Hannover Medical School, 30624 Hannover, Germany

a r t i c l e

i n f o

Article history: Received 9 February 2012 Received in revised form 7 May 2012 Accepted 14 May 2012 Available online 19 May 2012 Keywords: Polysialyltransferase Fluorescent polysialyltransferase acceptor HPLC separation Polymer elongation Carbohydrate polymerase Polysialic acid

a b s t r a c t Polysialyltransferases (polySTs) play critical roles in diverse biological processes, including neural development, tumorigenesis, and bacterial pathogenesis. Although the bacterial enzymes are presumed to have evolved to provide molecular mimics of the host-specific polysialic acid, no analytical technique is currently available to facilitate a direct comparison of the bacterial and vertebrate enzymes. Here we describe a new fluorescent acceptor, a 1,2-diamino-4,5-methylenedioxybenzene (DMB)-labeled trimer of a2,8-linked sialic acid (DMB–DP3), which primes both pro- and eukaryotic polySTs. High-performance liquid chromatography separation and fluorescence detection (HPLC–FD) of reaction products enabled the sensitive and quantitative detection of polyST activity, even using cell lysates as enzyme source, and revealed product profiles characteristic of each enzyme. Single product resolution afforded by this assay system revealed mechanistic insights into a kinetic lag phase exhibited by the polyST from Neisseria meningitidis serogroup B during chain elongation. DMB–DP3 is the first fluorescent acceptor shown to prime the mammalian polySTs. Moreover, product profiles obtained for the two murine polySTs provided direct biochemical evidence for enzymatic properties that had, until now, only been inferred from the analysis of biological samples. With DMB–DP3, we introduce a universal acceptor that provides an easy, fast, and reliable system for the comprehensive mechanistic and comparative analysis of polySTs. Ó 2012 Elsevier Inc. All rights reserved.

The term polysialic acid describes the series of linear homopolymers formed by the negatively charged nine-carbon sugar sialic acid (Sia).2 Different polysialic acid structures are defined by the chain length and the glycosidic linkage between residues. Several pathogenic bacteria synthesize a lipid-anchored capsule composed of the a2,8-, a2,9-, or alternating a2,8/a2,9-linked polysialic acid (Table 1). In vertebrates, exclusively the a2,8-linked polymer is synthesized as a posttranslational modification of a few specific protein scaffolds [1,2]. The enzymes responsible for biosynthesis of polysialic acid are known as polysialyltransferases (polySTs). These template-independent polymerases catalyze the successive addition of Sia from ⇑ Corresponding author. Fax: +49 511 532 8801. E-mail address: [email protected] (R. Gerardy-Schahn). These authors contributed equally to this work. Abbreviations used: Sia, sialic acid; polyST, polysialyltransferase; NCAM, neural cell adhesion molecule; FCHASE, 6-(fluorescein-5-carboxamido)hexanoic acid Nhydroxysuccimidyl ester; HPLC–FD, high-performance liquid chromatography with fluorescence detection; NmB, Neisseria meningitidis serogroup B; NmC, N. meningitidis serogroup C; DP3, trimeric a2,8-linked Sia; DMB, 1,2-diamino-4,5-methylenedioxybenzene; TFA, trifluoroacetic acid; UV, ultraviolet; DHB, 2,5-dihydroxybenzoic acid; MALDI–TOF MS, matrix-assisted laser desorption/ionization time-of-flight mass spectrometry; PCR, polymerase chain reaction; CA, colominic acid; DP, degree of polymerization. 1 2

0003-2697/$ - see front matter Ó 2012 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ab.2012.05.011

the activated sugar, CMP–Sia, onto the nonreducing end of a growing polysialic acid chain. Despite catalyzing the same polymerization reaction, the vertebrate and bacterial polySTs differ in the target of polysialylation (proteins and lipids, respectively) and do not share any sequence similarity. The sequence-based classification of carbohydrate-active enzymes (CAZy; see Ref. [3]) groups the eukaryotic polySTs with other sialyltransferases into the family GT 29, whereas the bacterial polySTs form their own family, GT 38 (for a review, see Ref. [4]). Currently, there is no method available that can analyze both groups of enzymes to provide further comparison of the polysialylation activities and their underlying mechanisms. The bacterial polySTs are localized on the cytoplasmic side of the inner membrane and are thought to associate with several transmembrane proteins to form the capsule synthesis and export complex [5]. Although the final capsular polysialic acid is anchored in the membrane via a phosphodiester-linked 1,2-dipalmitoyl glycerol [6], it is uncertain which molecule forms the initial priming acceptor for the bacterial enzymes [7]. Studies carried out in vitro have determined that the minimal glycoconjugate acceptors require the presence of di- or trisialylated oligosaccharides [8–10], but when primed with the free sugar, a trimer of sialic acid (DP3) was the minimum acceptor [11].

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Colominic acid was purchased from Sigma–Aldrich (Taufkirchen, Germany).

Table 1 Polysialic acid encapsulated bacteria. Bacterial strain

Linkage

Reference(s)

Escherichia coli K1 Neisseria meningitidis serogroup B Neisseria meningitidis serogroup C Escherichia coli K92

a2,8

[48,49] [50] [50] [51]

a2,9 a2,8/a2,9

In vertebrates, two polySTs (ST8SiaII and ST8SiaIV) are known to synthesize a2,8-linked polysialic acid. These enzymes predominantly recognize and polysialylate the neural cell adhesion molecule (NCAM) [1]. However, more recent studies also identified that SynCAM1 [12], PCAM [13], neuropilin-2 [14], CD36 [15], and the a-subunit of the voltage-gated sodium channel [16] can be polysialylated in certain physiological contexts. The small number of targets of polysialylation strongly suggests that polysialylation in vertebrates is a protein-specific process requiring an initial interaction between the polyST and the protein acceptor [17,18]. Chain length and dispersity of polysialic acid are likely to be critical determinants of its physiological roles in cell signaling, adhesion, and host–pathogen interactions [19]; however, relatively little is known of the molecular mechanisms governing initiation, elongation, and termination of polysialic acid products. Such mechanistic investigations require sensitive and accurate methods that provide quantitative information and single product resolution. Although radioactivity-based assays enable the sensitive and accurate quantification of incorporated Sia, they typically lack the resolution to distinguish individual product lengths. A major advance in bacterial polyST characterization was achieved with the introduction of a fluorescent acceptor, the ganglioside analogue GT3– FCHASE [6-(fluorescein-5-carboxamido)hexanoicacid N-hydroxysuccimidylester], combined with high-performance liquid chromatography with fluorescence detection (HPLC–FD) analysis of reaction products [20–22]. This acceptor has been shown to prime the polySTs from Escherichia coli K1 [10], E. coli K92 [21], Neisseria meningitidis serogroup B (NmB) [11], and N. meningitidis serogroup C (NmC) [23] and has led to the synthesis and analysis of other ganglioside-based fluorescent acceptors, including a trisialyllactosyl– BODIPY (boron dipyromethene) [23] and GD3–FCHASE [10]. Although these fluorescent acceptors have contributed mechanistic insights into bacterial polyST activity, the structures are relatively inaccessible due to multistep chemoenzymatic syntheses and have not been shown to prime polySTs of eukaryotic origin. Here we introduce a new non-ganglioside fluorescent acceptor consisting of a trimer of a2,8-linked Sia (DP3) directly conjugated to a 1,2-diamino-4,5-methylenedioxybenzene (DMB) label. This fluorescent acceptor is synthesized in a single step from commercially available reagents and purified to homogeneity by anion exchange chromatography. We establish that the acceptor is applicable to the quantitative and high-resolution analysis of mammalian and bacterial polySTs using either purified enzyme or cell-free extracts as enzyme source. A comparative analysis carried out with a panel of six polySTs highlighted diverse mechanisms of chain elongation and consequently dramatic differences in the length distribution of products. Altogether, the DMB–DP3 acceptor and HPLC–FD analysis provides a path to the comprehensive mechanistic characterization of diverse polySTs.

Materials and methods Materials CMP–Sia and DP3 were purchased from Nacalai Tesque (Kyoto, Japan). DMB was purchased from Dojindo (Kumamoto, Japan).

Synthesis and purification of DMB–DP3 For labeling DP3 with the fluorescent dye, DMB, reaction protocols established for the DMB labeling of oligo- and polysialic acid [24,25] were followed with minor modifications. Briefly, DP3 was dissolved at 10 mg ml1 in 20 mM DMB with 1 M b-mercaptoethanol and 40 mM sodium dithionite. The DP3 solution was then mixed with an equal volume of ice-cold 40 mM trifluoroacetic acid (TFA) and incubated for 48 h at 4 °C. The reaction was stopped by the addition of one-fifth reaction volume of 200 mM sodium hydroxide. Separation of 500-ll reaction aliquots was achieved by preparative anion exchange chromatography on a MonoQ 10/ 100 column (Amersham Biosciences). Chromatography was performed at 4 ml min1 with 10 mM Tris–HCl (pH 8.0) (M1) and with 1 M NaCl and 10 mM Tris–HCl (pH 8.0) (M2). After washing the column with 100% M1 for 5 min, the elution of DMB–DP3 was achieved with a linear gradient from 0 to 25% M2 over 20 min. The elution profile was monitored at 214 nm using an ultraviolet (UV) detector (SPD-20AV) and DMB fluorescence (excitation 373 nm/emission 456 nm) monitored with a fluorescence detector (RF-10A XL). A single fraction containing the DMB–DP3 was collected between 21 and 23 min. Fractions from several runs were pooled and freeze-dried. The dried material was dissolved in 4 ml of water, and salt was removed by extensive dialysis (1000 molecular weight cutoff [MWCO], ZelluTrans V-series, Roth) against water. The desalted DMB–DP3 was freeze-dried, weighed, and dissolved at 50 mM in water. DMB–DP3 samples prepared in this way were stable for more than 12 months when stored at 80 °C, with no detectable degradation or loss of fluorescence. Mass spectrometric analysis For matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI–TOF MS) sample preparation, 0.7 ll of the purified product (0.7 mM DMB–DP3) was mixed on the target plate with 0.7 ll of 2,5-dihydroxybenzoic acid (DHB, 5 mg ml1 in 50% acetonitrile with 0.1% TFA, Bruker Daltonics, Billerica, MA, USA) and subsequently air-dried. MALDI–TOF MS was carried out on a Voyager-DE PRO mass spectrometer (Applied Biosystems, Foster City, CA, USA) controlled by Voyager Control Panel 5.10.2 software. Spectra were acquired in negative ion reflection mode, averaging 1000 laser shots per MALDI–TOF spectrum. The spectra were displayed with Data Explorer 4.8 software (Applied Biosystems) without further processing. Construction of bacterial polyST expression vectors All DNA isolations were done using a Machery–Nagel NucleoSpin Plasmid or Extract kit. Enzymes were purchased from New England Biolabs, and all steps were carried out according to the manufacturer’s instructions. The pET32a expression vector (Novagen) was modified by replacing the thioredoxin tag with a short sequence encoding the Strep tag-II [26]. The full-length polyST genes were polymerase chain reaction (PCR) amplified using Phusion polymerase, according to the manufacturer’s instructions, and the following primer pairs: NmB–polyST, 50 -TGGCTGATATCGGATCCCTAAAG and 30 -CGAGTGCGGCCGCTCTATCTC; NmC–polyST, 50 -CCCTGGATATCGGATCCTTGCAGAAAATAAGAAAAGCTC and 30 -ACCCTGCGGCCGCTTG GTTACAAAGGCTATATTTA; K1–polyST, 50 -TGGCTGATATCGGATCCATGATATTTGATGCTAGTTTAAAG and 30 -ATATACTCGAGTGCGGCC GCCGAGTGCGGCCGCTCTATCTC; and K92–polyST, 50 -GCTGGGATATCGGATCCATATTTGATGCTAGTTTAAA GAAG and 30 -ACCCTGCG GCCGCCTCCCCCAAGAAAATCCTTTTA. PCR products were digested

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with BamHI and XhoI and were ligated into the similarly digested expression vector. The sequence of all constructs was confirmed by sequencing the cloned gene and flanking regions. Obtained vectors drive the inducible expression of full-length polySTs lacking only Met1 and flanked by an N-terminal polypeptide (MDSWSHP QFEKGALDIGS) that includes a Strep tag-II and a C-terminal polypeptide (AAALEHHHHHH) containing a 6His tag. All DNA manipulations were carried out according to the Deutsche Gentechnikgesetz (German Genetic Engineering Act).

gradient (shown in Fig. 3A in Results) consisting of a curved gradient from 0 to 26% M2 over 7 min followed by a linear gradient from 26 to 50% M2 over 22 min. The curved portion of the gradient is a number 3 curve (LC Solution, Shimadzu) and is described by

Preparation of recombinant polySTs

Results

The maltose binding protein-tagged NmB–polyST and the murine polySTs ST8SiaII and ST8SiaIV carrying 6His tags and N-term truncations of 56 and 25 amino acids, respectively, were expressed and purified as described previously [11,12]. The four bacterial polySTs (E. coli K1, E. coli K92, NmB, and NmC), with N-terminal Strep tag-II and C-terminal 6His tag, were expressed in E. coli BL21gold (DE3) (Stratagene). Bacteria were cultivated in Power Broth medium (Athena) containing 200 lg ml1 carbenicillin at 37 °C until an OD600 of 1.9 was reached. Cultures were rapidly cooled in an ice bath, supplemented with 0.5 mM isopropyl b-D1-thiogalactopyranoside (IPTG) to induce protein expression, and then allowed to grow at 15 °C for 20 h. Cultures typically reached an OD600 of 7.5. Cells were harvested by centrifugation at 6000g for 10 min, and pellets were stored at 20 °C prior to analysis. Cell pellets were resuspended in lysis buffer (50 mM Tris–HCl [pH 8.0], 5% glycerol, and 25 mM KCl) to a calculated OD600 of approximately 25, supplemented with 50 lg ml1 DNase I and 100 lg ml1 lysozyme, incubated for 15 min on ice, and subjected to two cycles of freeze/thawing. Obtained lysates were centrifuged at 16,000g at 4 °C for 20 min, and supernatants were used directly in activity assays.

Determination of minimal DMB-labeled acceptor structure

where t is time (in minutes).

To determine the minimal DMB-labeled acceptor structure used by polySTs, a panel of two mammalian and four bacterial polySTs was tested with DMB-labeled colominic acid (DMB–CA). CA is the partially hydrolyzed, purified capsular polysaccharide from E. coli K1. The DMB–CA sample contains the complete range of potential a2,8-linked acceptor structures from the DMB-labeled sialylmonomer (DMB–DP1) to long DMB-labeled polysialic acid chains (Fig. 1A). Reactions were carried out with a 200-fold molar excess of donor sugar (CMP–Sia) with respect to the acceptor and were allowed to proceed to completion. HPLC analysis of the reaction products showed that none of the enzymes transferred Sia onto DMB–DP1 or DMB–DP2 (Fig. 1B). In accord with previous studies indicating that CA is a poor acceptor for the mammalian polySTs and the E. coli K92 polyST [9,28–30], we saw little modification of the DMB–CA acceptor by these enzymes (data not shown). In contrast, DMB–DP3 as well as larger DMB oligo- and polymers were effectively elongated by the a2,8-polySTs from NmB and E. coli K1 and by the a2,9-polySTs from NmC (Fig. 1B). Based on

A

DMB-Labelled Colominic Acid

DP6

DP5

DP2 DP1

Fluorescence intensity

3 2 Retention time (min)

25

20

4

PolyST Reactions

NmB-polyST

Fluorescence intensity

B

1

15

DP4

10

0 12 3 4 5

Testing of polySTs To determine the activity of polySTs, the recombinant enzymes were assayed at 25 °C in 10- to 200-ll volumes containing 50 mM Tris–HCl (pH 8.0), 25 mM KCl, 20 mM MgCl2, 5% glycerol, 0.01 to 1 mM CMP–Sia, and 5 to 200 lM DMB–DP3. The reaction buffer for testing of the mammalian polySTs was 10 mM sodium cacodylate (pH 6.7) supplemented with 10 mM MnCl2. Reactions were started with the addition of enzyme to give a final protein concentration of 150 lg ml1 for bacterial lysates or 25 to 50 lg ml1 for the purified NmB–polyST or mammalian polySTs. Reactions were stopped by a 10-fold dilution of the sample in 100 mM Tris–HCl (pH 8.0) and 20 mM ethylenediaminetetraacetic acid (EDTA), followed by 10 min at 50 °C. Samples were centrifuged at 20,000g for 20 min or filtered through 0.22-lm filters (Millipore) prior to analysis by HPLC. HPLC analysis was carried out on a UFLC-RX system (Shimadzu) fitted with a UV detector (SPD-20AV) and a fluorescence detector (RF-10A XL). Between 10 and 50 ll of stopped reaction sample, corresponding to 5 to 20 pmol of acceptor, was applied to a CarboPac PA-100 column (Dionex, Sunnyvale, CA, USA) for separation. Samples were separated according to established methods [25,27] with minor modifications as follows. Chromatography was conducted at 0.6 ml min1 with a column temperature of 50 °C, with the mobile phases consisting of 20 mM NaNO3 (M1) and 1 M NaNO3 (M2). Products were separated using an elution

ð1Þ

DP3

Protein concentration determinations were carried out with the BCA (bisinchoninic acid) assay (Pierce) according to the manufacturer’s instructions.

26  ðe3t=7  1Þ ; ðe3  1Þ

Fluorescence intensity

Protein determinations

%M2 ¼

NmC-polyST

K1-polyST 1

3 2 Retention time (min)

4

Fig.1. Determination of minimal DMB-labeled acceptor structure. (A) HPLC separation of DMB-labeled colominic acid shows the presence of the full range of potential labeled acceptor structures from DP1 to long polysialic acid chains. (B) Reaction products of bacterial polySTs primed with the DMB-labeled colominic acid show that the labeled DP1 and DP2 are not modified by the enzymes. However, the labeled DP3 and longer oligomers were effectively elongated by the NmB, NmC, and E. coli K1 polySTs.

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product profiles indicates that the oligo- and polysialic acid reaction products have a higher affinity for the enzyme than the initial DMB–DP3 acceptor. At the 10-min reaction time point, degrees of polymerization (DPs) 4 to 7 are more abundant than DMB–DP3, whereas by 20 min and in all subsequent reaction time points, DPs 4 to 7 are considerably less abundant than DMB–DP3 and long polymeric products eluting at retention times of 18 to 22.5 min accumulate. The results show that the NmB–polyST has a higher reaction rate with the oligo- and polysialic acid products than with DMB–DP3 and indicate an increasing affinity for the extended reaction products. The elution gradient (Fig. 3A) used in this study enabled the resolution of individual peaks for products up to approximately DP53. By extrapolating from the retention time of resolved peaks over the linear portion of the elution curve, we were able to estimate the chain lengths of unresolved products (Fig. 3C). The longest products synthesized by the NmB–polyST were estimated to range between DP110 and DP160. These chain lengths agree well with the size of polysialic acid isolated directly from Neisseria or E. coli K1 cultures that range in length between DP130 and DP200 [31–33].

these data, we decided to synthesize and purify DMB–DP3 as acceptor substrate for the development of an HPLC-based polyST assay. Synthesis and purification of DMB–DP3 The DMB-labeling reaction was carried out under conditions previously optimized by Inoue and coworkers for labeling of CA [25]. To minimize hydrolysis of glycosidic bonds under the acidic conditions, the reaction was maintained at 4 °C. Reactions were terminated by the addition of NaOH after 48 h, and products were separated by anion exchange chromatography as described in Materials and Methods. Elution profiles were monitored via absorption at 214 nm and DMB fluorescence, which enabled progress of the labeling and hydrolysis reactions to be followed. The retention time of unmodified DP3 enabled a clear separation from the mixture of hydrolysis products and reagents, which eluted with a retention time of up to 18 min, and the desired product DMB–DP3, which eluted at a retention time of 21 to 22.5 min (Fig. 2A). Reanalysis of the collected DMB–DP3 peak (after concentration and desalting) demonstrated purity of the product (Fig. 2A). The synthesis and purification produced 10.4 mg of DMB–DP3 in an overall yield of 38%. To confirm the identity of the collected peak with the expected DMB–DP3 structure (Fig. 2B), MS analysis was performed. MALDI–TOF spectra (Fig. 2C) showed strong signals for masses corresponding to DMB–DP3, the first and second sodium salt, and the first (-H2O) and second (-2H2O) lactonized forms. Lactonization of DP3 is a reversible modification resulting from acidic conditions in the MALDI matrix [24] and is not expected to occur at neutral pH.

Quantitative evaluation of NmB–polyST activity To extend the use of the newly established HPLC assay toward quantitative analysis of polyST activity, it was necessary to establish a method suited to calculate the number of glycosidic bonds formed at each measured time point. Products with DP > 50 are insufficiently resolved to distinguish peaks for the individual DPs; therefore, the length of synthesized polymers was limited to DP < 50 by using up to a 15-fold molar excess of donor sugar with respect to DMB–DP3. Reactions were carried out in duplicate using 20 lM DMB–DP3, 250 lM donor sugar, and three enzyme concentrations. An example chromatogram obtained after 60 min of incubation with 150 lg ml1 NmB–polyST is shown in Fig. 4A. The amount of each DP in the product profile was determined by integration of the HPLC profiles, with the boundary of each DP being

Product profiles of NmB–polyST reaction with DMB–DP3 To observe the elongation of DMB–DP3 by the purified NmB– polyST, a series of reaction time points were analyzed by HPLC. The DMB–DP3 acceptor (Fig. 3A) was gradually extended to yield long polysialic acid products (Fig. 3B). Detailed analysis of the

A

30 20

214 nm Fluorescence

DMB-DP3 (theoretical mass: 1007.33)

B

100 DP3 (sialic acid trimer)

OH HO

50

OH

DP3 labeling reaction

200

50 100 0 30

0 100 Purified DMB-DP3

20

50 10

HO

O

HO

O OH

HO

O

N

HO

OH OH

[M -H -H2O]988.32

100

0 10 15 20 25 Retention time (min)

30

35

N

OH

AcHN

C

HO

80 [M -H]1006.29

60 40

[M -2H -H2O +Na]1010.31

[M -H -2H2O]970.27

[M -2H +Na]1028.32 [M -3H +2Na]1050.31

20

5

O

COOH

AcHN

% Intensity

0 100

Fluorescence intensity

A214

0 300

0 0

O

AcHN

10

O

COOH

0 960

980

1000

1020 Mass (m/z)

1040

1060

Fig.2. Purification and characterization of DMB–DP3. Samples from the synthesis and purification of DMB–DP3 were separated on a preparative MonoQ column and monitored via UV absorbance and fluorescence of the DMB label (excitation 373 nm/emission 456 nm). (A) Unlabeled DP3 can be observed at a retention time of 30 min in the 214-nm channel and does not show any detectable fluorescence. Separation of the labeling reaction shows the consumption of DP3 and the appearance of fluorescent species. The most prominent fluorescent product at a retention time of 22 min corresponds to the desired DMB–DP3 product and was collected for further analysis. The purity of the collected DMB–DP3 was confirmed by reanalysis on the MonoQ column. (B) Chemical structure and theoretical mass of the DMB–DP3 acceptor substrate. (C) The identity of the purified product was confirmed by MALDI–TOF MS analysis. The measured value and the expected mass of each species are indicated above each peak.

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A

75

Elution Gradient

50 25

0

5.0

10.0

15.0

20.0

% 1M NaNO3

Fluorescence intensity

100 DMB-DP3

25.0

B 5 min

Fluorescence intensity

10 min

20 min

30 min

60 min

16 hr 0

5.0

10.0

15.0

20.0

25.0

Retention time (min)

C

Degree of Polymerisation 3 0

10

20

5.0

40

30 10.0

50

60 15.0

70

80 90 100

120 20.0

140

160 180 25.0

Retention time (min) Fig.3. Analysis of NmB–polyST activity with the DMB–DP3 acceptor. The enzymatic synthesis of polysialic acid onto the DMB–DP3 acceptor was followed by the separation of reaction samples on a CarboPac-PA100 column (Dionex). (A) The synthesized DMB–DP3 acceptor elutes as a single peak with a retention time of approximately 2.5 min using the displayed elution gradient. (B) The analysis of successive reaction time points shows the gradual elongation of the acceptor structure, resulting in the formation of long polysialic acid products. (C) The DP ruler indicates the retention times of resolved peaks in black and the estimated lengths of unresolved chain lengths in gray.

defined by a vertical line from the peak trough to the baseline (Fig. 4A). Given the quantity of each DP, the number of glycosidic bonds formed is given by

formed glycosidic bonds ¼

50 X

xn  ðn  3Þ;

ð2Þ

n¼4

where xn is the molar amount of the nth DP. Importantly, the progress curves show that the NmB–polyST does not follow a typical enzymatic reaction kinetic (Fig. 4B). The polymerization reaction does not proceed with an initial maximum velocity; rather, the maximum reaction rate occurs after consumption of 50 to 100 lM CMP–Sia. Such a kinetic lag phase is a common feature of carbohydrate polymerases that progressively modify their acceptor substrates [34]. The obtained data are in perfect agreement with our earlier observations. Using a continuous spectrophotometric assay and unlabeled oligosialic acid acceptors (DP3–DP5), an initial lag phase was observed with the NmB–polyST [11]. Detection of bacterial polyST activity in crude cell extracts Being membrane-associated proteins, the bacterial polySTs have typically proven to be difficult to overexpress and purify. Therefore, polyST activity is often investigated in bacterial lysates

using radiochemical assays, which are sensitive, specific, and robust [7,9,35] but do not provide information on the length or abundance of specific reaction products. Aiming at providing an assay of similar robustness but more sophisticated in terms of product analysis, we tested the newly established assay with four bacterial polySTs (from E. coli K1, E. coli K92, NmB, and NmC) exhibiting different linkage specificities (see Table 1). The enzymes were cloned with only short N-terminal (Strep tag-II) and C-terminal (6His) tags and were overexpressed in E. coli BL21 (DE3). Cell lysates were prepared, debris was removed, and polyST activities were tested directly in the cell-free extracts by the addition of reaction buffer, donor sugar, and DMB–DP3 acceptor. The stability of the DMB–DP3 acceptor was established by incubation with mock lysates (containing no enzyme). For more than 24 h, no modification of the DMB–DP3 peak could be observed in the presence of the E. coli lysate independent of the absence or presence of CMP–Sia (data not shown). However, DMB–DP3 was elongated in all preparations containing the recombinant expressed bacterial polySTs. Each of the polySTs exhibited characteristic patterns of acceptor elongation. As already described for the purified NmB–polyST, the products of the NmB–polyST in the cell-free extract indicated continuous elongation of the DMB–DP3 acceptor from short to long chains. In contrast, the NmC–polyST, which forms a2,9 linkages, displayed a biphasic mode of acceptor elongation. Polymers with up to DP15 (DMB–a2,8Sia3-a2,9Sia12) were synthesized in an

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A

0 0.0

5.0

DP38+

20

DP25

DP20

40

DP30

DP7

DP1 5

60 DP3

Fluorescence intensity

Peak Area Determination

10.0

15.0

20.0

Retention time (min)

B

Progress Curves

Glycosidic bonds formed (µM)

250 200 150

the bacterial enzymes except for the reaction buffer, which was optimized for the mammalian polySTs. Despite the high acceptor specificity of the mammalian polySTs and the lack of protein and N-glycan components present in the natural acceptor, both enzymes catalyzed the transfer of Sia onto the DMB–DP3 acceptor. Remarkably, very different patterns of chain elongation were observed. ST8SiaII efficiently used DMB–DP3 (only 1.2% of the acceptor remained after overnight incubation) to produce oligomers with an approximately normal distribution around DP7 (Fig. 5B). This absence of products with DP > 20 indicated that the enzyme has little affinity for longer polysialic acid chains. In stark contrast, the product profile of the ST8SiaIV reaction showed that more than 10% of the DMB–DP3 remained unmodified and chains of DP > 60 represent the major product (Fig. 5B). Although the synthesized oligosialic acid chains (DP4–DP10) remain prominent in the final products, there was no significant accumulation of other intermediates on the path to long chains. The observed product pattern demonstrates the increasing affinity of ST8SiaIV for longer chains.

150 µg/ml

100

100 µg/ml 50 µg/ml

50 0 0

10

20

30

40

50

60

70

80

90 100

110 120 130

Reaction time (min)

Fig.4. Quantification of NmB–polyST reactions. For each HPLC chromatogram, the relative abundance of each synthesized DP was estimated by integration of the HPLC profile and normalization to the total area under the curve using LC Solution software (Shimadzu). (A) Example chromatogram showing 60-min time point of the 150-lg ml1 NmB–polyST reaction. Vertical gray lines indicate the peak divisions set by LC Solution to determine the area for each product. (B) The number of glycosidic bonds formed (sialic acid transfers) at each time point was calculated and plotted against the reaction time for three different NmB–polyST concentrations. Data points are from a single determination representative of two independent experiments.

initial phase, followed by the slow accumulation of long a2,9 polymers without the apparent accumulation of intermediates (Fig. 5A). In addition, the long reaction products of the NmC–polyST display a much broader distribution than the products formed by NmB–polyST. Together, the data indicate that the enzymes from N. meningitides serogroups B and C exhibit significantly different mechanisms of elongation under these assay conditions. The E. coli K1 polyST catalyzes formation of a2,8 linkages; however, it did not form long polysialic acid products on the DMB–DP3 acceptor (Fig. 5A). The most abundant products were oligosialic acid of approximately DP8, whereas products with DP > 30 were not detectable. This finding confirms a previous study. Using FCHASE–aminophenylglycoside as acceptor, Willis and coworkers [10] also demonstrated that the E. coli K1–polyST only synthesizes oligosialic acid. In contrast to the other enzymes, the E. coli K92– polyST showed very little activity. After overnight incubation, the DMB–DP3 acceptor remained mostly unmodified; however, chains of up to DP9 could be detected (Fig. 5A). Analysis of mammalian polySTs The mammalian polySTs are responsible for synthesis of polysialic acid onto specific glycoprotein acceptors. Whereas fluorescent acceptors have been used for the testing of bacterial polySTs (as described above), no such high-resolution assay system has been reported for the mammalian polySTs. Here we investigated whether DMB–DP3 can also be applied to analyze mammalian polySTs. Purified recombinant enzymes carrying N-terminal 6His tags and truncations of the N-terminal transmembrane domain [12] were assayed under conditions identical to those used for

Discussion To obtain a highly pure and easy-to-handle fluorescent acceptor for the fast and reliable testing of polySTs, we have investigated DMB-labeled oligosialic acid acceptors and found the DMB–DP3 acceptor to be the smallest structure that is recognized and elongated by a range of polySTs. The simple one-step synthesis and chromatographic purification of DMB–DP3 is a considerable advantage over the previously described fluorescent ganglioside analogues that are synthesized via a multistep chemoenzymatic route and require considerable effort to yield milligram amounts of the homogeneous compounds [10,20,22,23,36]. Although fluorescent ganglioside analogues have been used for testing bacterial polySTs [10,11,21,23], DMB–DP3 is the first fluorescent acceptor shown to prime the eukaryotic polySTs and, thus, enable the comparative testing of these different classes of enzymes. The DMB–DP3 acceptor was first evaluated using a purified and well-characterized NmB–polyST construct [11]. With a 200-fold molar excess of CMP–Sia, long polysialic acid chains similar to those observed in earlier studies with nonlabeled oligosialic acid acceptors [11] were synthesized on the DMB–DP3 acceptor. Interestingly, time lapse recording of the reaction progress indicated that NmB–polyST products were skewed toward longer polysialic acid chains. The effect is best visible up to the 20-min time point in Fig. 3B, where significant amounts of the starting material remain, and the consumption of newly synthesized longer DPs is favored. The quantitative estimation of the NmB–polyST reaction revealed that this effect corresponds to a kinetic lag phase, indicating that the accelerating initial reaction rate is caused by the enzyme’s increasing affinity for its reaction products. Kinetic lag phases have been described for other polymerizing glycosyltransferases, including the E. coli K92 polyST [21], the hyaluronan synthase from Pasteurella multocida [37], and the capsular polysaccharide type 3 synthase from Streptococcus pneumonia [38]. Furthermore, a similar mechanism was recently shown to underlie the lag phase of the model carbohydrate polymerase, GlfT2, a galactofuranose polymerase involved in Mycobacterium tuberculosis cell wall biosynthesis [34]. Importantly, the kinetic lag phase was also observed when the NmB–polyST was tested with the unlabeled oligosialic acid (DP3, DP4, and DP5) acceptors in a continuous spectrophotometric assay [11], indicating that the effect cannot be attributed to the fluorescently labeled acceptor. A major advantage of the current assay is broad applicability, high reproducibility, and the complete absence of background signals, also when carried out in complex biological samples. To interrogate the mechanism of polyST chain elongation in near-native

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Bacterial PolySTs 40

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Fig.5. Distribution of polysialic acid synthesized by different bacterial and mammalian polySTs. (A) The activity of four bacterial polySTs was measured directly in cell-free extracts. Reactions containing 5.32 lM DMB–DP3 and 1 mM CMP–Sia were started by the addition of cell-free extract to the final protein concentration of approximately 150 lg ml1. Reaction samples containing 5.32 pM acceptor were analyzed by HPLC. HPLC chromatograms of the final product distribution for all of the bacterial polySTs are shown. (B) The purified mammalian polySTs (ST8SiaII and ST8SiaIV) were assayed with the same concentration of acceptor and donor substrates. The HPLC chromatograms of the product distribution after overnight reaction are displayed.

conditions, we tested four bacterial polySTs directly in cell-free extracts. The enzymes were expressed with only short epitope tags to limit interference with the natural localization of the proteins on the cytoplasmic side of the inner membrane [21,35,39]. The analysis illustrated that each polyST synthesized a characteristic pattern of products, indicating differences in the mechanisms of chain elongation used by each enzyme. Whereas the patterns displayed with the polySTs from NmB, NmC, and E. coli K1 are consistent with previously described reaction profiles [10,11,23], the profile displayed with polyST from E. coli K92 deviates. Although we could observe transfers, DMB–DP3 seems to be a suboptimal acceptor for this enzyme. Under similar assay conditions and with GT3– FCHASE as an acceptor, Vionnet and Vann [21] detected formation of long polymers already after very short incubation times. The glycan portion of GT3–FCHASE is trisialyllactose; this considerably larger structure may provide an improved geometry for the acceptor binding site in the E. coli K92 enzyme. Alternatively, the extended hydrophobic linker and fluorescein label may interact with hydrophobic surfaces of the K92–polyST, effectively tethering the acceptor to the enzyme. Indeed, such a tethering effect has

been demonstrated with the model enzyme, GlfT2, where hydrophobic linkers of increasing length increased the number of transfers onto the acceptor [40]. Nevertheless, DMB–DP3 was elongated by the E. coli K92–polyST, and the low activity effectively demonstrates the sensitivity afforded by the zero-background and femtomolar detection of the DMB label [25], making the assay system attractive, for instance, to detect activity in native enzyme sources. The DMB–DP3 acceptor is the first fluorescent acceptor shown to prime the mammalian polySTs, and (surpassing our expectations) the obtained product profiles exactly reflect the enzymatic features that have been attributed to the murine polySTs based on indirect evidence obtained by analysis of biological samples [41–44]. In more detail, using mouse strains with genetically induced defects in expression of the polySTs, it became obvious that ST8SiaII has a much higher potential to initiate the transfer of polysialic acid onto the major carrier protein NCAM. Approximately 45% of the NCAM pool remained polysialic acid free in ST8SiaII knockout mice, whereas depletion of ST8SiaIV allowed only 10% of the total NCAM pool to escape polysialylation [42].

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However, ST8SiaIV knockout significantly affected the production of long polysialic acid chains [41] and, thus, confirmed in vitro studies demonstrating that ST8SiaII and ST8SiaIV in combination synthesize more polysialic acid on NCAM than does either of the enzymes in a separate reaction [45]. The data from the DMB–DP3 acceptor confirm and strengthen the models of murine polyST activity that were derived from these earlier studies. The ST8SiaII effectively elongates short primers and produces oligomeric products, whereas ST8SiaIV has little affinity for short chains but readily produces long polysialic acid given an oligomeric acceptor. The congruence of the described data allows us to predict that the DMB–DP3 acceptor will gain an important position in the analysis of mammalian polySTs. For instance, this system should be highly suitable to evaluate the function of structural elements that distinguish the vertebrate polySTs from other closely related members of the sialyltransferase family [46,47]. In conclusion, the DMB-DP3 acceptor is accessible via a simple one-step synthesis and provides a robust and highly sensitive assay format for testing of polySTs from both pro- and eukaryotic origin. The quantitative information gained in terms of both the rate of sugar transfer, and the distribution of products throughout the reaction course provide a comprehensive description of the polyST reaction which will continue to give new mechanistic insights into this important class of enzyme. Acknowledgments Financial support for this work was provided by the Deutsche Forschungsgemeinschaft (DFG) in the framework of DFG Research Unit 548 (Ge801/10-1). We thank David Schwarzer for many helpful discussions and Martina Mühlenhoff for critical comments on the manuscript. References [1] H. Hildebrandt, M. Mühlenhoff, R. Gerardy-Schahn, Polysialylation of NCAM, in: V. Berezin (Ed.), Structure and Function of the Neural Cell Adhesion Molecule NCAM, Springer, New York, 2010, pp. 95–109. [2] U. Rutishauser, Polysialic acid in the plasticity of the developing and adult vertebrate nervous system, Nat. Rev. Neurosci. 9 (2008) 26–35. [3] B.L. Cantarel, P.M. Coutinho, C. Rancurel, T. Bernard, V. Lombard, B. Henrissat, The Carbohydrate-Active enZymes database (CAZy): an expert resource for glycogenomics, Nucleic Acids Res. 37 (2009) D233–D238. [4] M. Audry, C. Jeanneau, A. Imberty, A. Harduin-Lepers, P. Delannoy, C. Breton, Current trends in the structure–activity relationships of sialyltransferases, Glycobiology 21 (2011) 716–726. [5] S.M. Steenbergen, E.R. Vimr, Biosynthesis of the Escherichia coli K1 group 2 polysialic acid capsule occurs within a protected cytoplasmic compartment, Mol. Microbiol. 68 (2008) 1252–1267. [6] Y.L. Tzeng, A.K. Datta, C.A. Strole, M.A. Lobritz, R.W. Carlson, D.S. Stephens, Translocation and surface expression of lipidated serogroup B capsular polysaccharide in Neisseria meningitidis, Infect. Immun. 73 (2005) 1491–1505. [7] C. Weisgerber, F.A. Troy, Biosynthesis of the polysialic acid capsule in Escherichia coli K1: the endogenous acceptor of polysialic acid is a membrane protein of 20 kDa, J. Biol. Chem. 265 (1990) 1578–1587. [8] J.W. Cho, F.A. Troy, Polysialic acid engineering: synthesis of polysialylated neoglycosphingolipids by using the polysialyltransferase from neuroinvasive Escherichia coli K1, Proc. Natl. Acad. Sci. USA 91 (1994) 11427–11431. [9] M.M. McGowen, J. Vionnet, W.F. Vann, Elongation of alternating a2,8/2,9 polysialic acid by the Escherichia coli K92 polysialyltransferase, Glycobiology 11 (2001) 613–620. [10] L.M. Willis, M. Gilbert, M.-F. Karwaski, M.-C. Blanchard, W.W. Wakarchuk, Characterization of the a-2,8-polysialyltransferase from Neisseria meningitidis with synthetic acceptors, and the development of a self-priming polysialyltransferase fusion enzyme, Glycobiology 18 (2008) 177–186. [11] F. Freiberger, H. Claus, A. Gunzel, I. Oltmann-Norden, J. Vionnet, M. Muhlenhoff, U. Vogel, W.F. Vann, R. Gerardy-Schahn, K. Stummeyer, Biochemical characterization of a Neisseria meningitidis polysialyltransferase reveals novel functional motifs in bacterial sialyltransferases, Mol. Microbiol. 65 (2007) 1258–1275. [12] S.P. Galuska, M. Rollenhagen, M. Kaup, K. Eggers, I. Oltmann-Norden, M. Schiff, M. Hartmann, B. Weinhold, H. Hildebrandt, R. Geyer, M. Mühlenhoff, H. Geyer, Synaptic cell adhesion molecule SynCAM 1 is a target for polysialylation in postnatal mouse brain, Proc. Natl. Acad. Sci. USA 107 (2010) 10250–10255.

[13] M. Langhauser, J. Ustinova, E. Rivera-Milla, D. Ivannikov, C. Seidl, C. Slomka, J. Finne, Y. Yoshihara, M. Bastmeyer, J. Bentrop, Ncam1a and Ncam1b—two carriers of polysialic acid with different functions in the developing zebrafish nervous system, Glycobiology 22 (2012) 196–209. [14] S. Curreli, Z. Arany, R. Gerardy-Schahn, D. Mann, N.M. Stamatos, Polysialylated neuropilin-2 is expressed on the surface of human dendritic cells and modulates dendritic cell–T lymphocyte interactions, J. Biol. Chem. 282 (2007) 30346–30356. [15] U. Yabe, C. Sato, T. Matsuda, K. Kitajima, Polysialic acid in human milk, J. Biol. Chem. 278 (2003) 13875–13880. [16] C. Zuber, P.M. Lackie, W.A. Catterall, J. Roth, Polysialic acid is associated with sodium channels and the neural cell adhesion molecule N-CAM in adult rat brain, J. Biol. Chem. 267 (1992) 9965–9971. [17] D.A. Foley, K.G. Swartzentruber, K.J. Colley, Identification of sequences in the polysialyltransferases ST8Sia II and ST8Sia IV that are required for the proteinspecific polysialylation of the neural cell adhesion molecule, NCAM, J. Biol. Chem. 284 (2009) 15505–15516. [18] D.A. Foley, K.G. Swartzentruber, M.G. Thompson, S.S. Mendiratta, K.J. Colley, Sequences from the first fibronectin type III repeat of the neural cell adhesion molecule allow O-glycan polysialylation of an adhesion molecule chimera, J. Biol. Chem. 285 (2010) 35056–35067. [19] D. Nakata, F.A. Troy, Degree of polymerization (DP) of polysialic acid (PolySia) on neural cell adhesion molecules (N-CAMs), J. Biol. Chem. 280 (2005) 38305– 38316. [20] O. Blixt, D. Vasiliu, K. Allin, N. Jacobsen, D. Warnock, N. Razi, J.C. Paulson, S. Bernatchez, M. Gilbert, W. Wakarchuk, Chemoenzymatic synthesis of 2azidoethyl-ganglio-oligosaccharides GD3, GT3, GM2, GD2, GT2, GM1, and GD1a, Carbohydr. Res. 340 (2005) 1963–1972. [21] J. Vionnet, W.F. Vann, Successive glycosyltransfer of sialic acid by Escherichia coli K92 polysialyltransferase in elongation of oligosialic acceptors, Glycobiology 17 (2007) 735–743. [22] W.W. Wakarchuk, A.-M. Cunningham, Capillary electrophoresis as an assay method for monitoring glycosyltransferase activity, in: P. Thibault, S. Honda (Eds.), Capillary Electrophoresis of Carbohydrates, Humana, Totowa, NJ, 2003, pp. 263–274. [23] D.C. Peterson, G. Arakere, J. Vionnet, P.C. McCarthy, W.F. Vann, Characterization and acceptor preference of a soluble Meningococcal group C polysialyltransferase, J. Bacteriol. 193 (2011) 1576–1582. [24] S.P. Galuska, R. Geyer, M. Mühlenhoff, H. Geyer, Characterization of oligo- and polysialic acids by MALDI–TOF–MS, Anal. Chem. 79 (2007) 7161–7169. [25] S. Inoue, S.-L. Lin, Y.C. Lee, Y. Inoue, An ultrasensitive chemical method for polysialic acid analysis, Glycobiology 11 (2001) 759–767. [26] T.G. Schmidt, A. Skerra, The Strep-tag system for one-step purification and high-affinity detection or capturing of proteins, Nat. Protoc. 2 (2007) 1528– 1535. [27] S. Inoue, Y. Inoue, Ultrasensitive analysis of sialic acids and oligo [+45 degree rule] polysialic acids by fluorometric high-performance liquid chromatography, in: Y.C. Lee, R.T. Lee (Eds.), Recognition of Carbohydrates in Biological Systems, part A: General Procedures, Academic Press, San Diego, 2003, pp. 543–560. [28] C.-F. Chao, H.-C. Chuang, S.-T. Chiou, T.-Y. Liu, On the biosynthesis of alternating a-2,9/a-2,8 heteropolymer of sialic acid catalyzed by the sialyltransferase of Escherichia coli Bos-12, J. Biol. Chem. 274 (1999) 18206– 18212. [29] K. Angata, M. Suzuki, J. McAuliffe, Y. Ding, O. Hindsgaul, M. Fukuda, Differential biosynthesis of polysialic acid on neural cell adhesion molecule (NCAM) and oligosaccharide acceptors by three distinct a2,8-sialyltransferases, ST8Sia IV (PST), ST8Sia II (STX), and ST8Sia III, J. Biol. Chem. 275 (2000) 18594–18601. [30] R.D. Mccoy, E.R. Vimr, F.A. Troy, CMP–NeuNAc–poly-a-2,8-sialosyl sialyltransferase and the biosynthesis of polysialosyl units in neural cell adhesion molecules, J. Biol. Chem. 260 (1985) 2695–2699. [31] B. Rode, C. Endres, C. Ran, F. Stahl, S. Beutel, C. Kasper, S. Galuska, R. Geyer, M. Mühlenhoff, R. Gerardy-Schahn, T. Scheper, Large-scale production and homogenous purification of long chain polysialic acids from E. coli K1, J. Biotechnol. 135 (2008) 202–209. [32] S. Pelkonen, J. Aalto, J. Finne, Differential activities of bacteriophage depolymerase on bacterial polysaccharide: binding is essential but degradation is inhibitory in phage infection of K1-defective Escherichia coli, J. Bacteriol. 174 (1992) 7757–7761. [33] R. Chen, J. John, B. Rode, B. Hitzmann, R. Gerardy-Schahn, C. Kasper, T. Scheper, Comparison of polysialic acid production in Escherichia coli K1 during batch cultivation and fed-batch cultivation applying two different control strategies, J. Biotechnol. 154 (2011) 222–229. [34] M.R. Levengood, R.A. Splain, L.L. Kiessling, Monitoring processivity and length control of a carbohydrate polymerase, J. Am. Chem. Soc. 133 (2011) 12758–12766. [35] I.K. Vijay, F.A. Troy, Properties of membrane-associated sialyltransferase of Escherichia coli, J. Biol. Chem. 250 (1975) 164–170. [36] S.L. Mosley, P.C. Rancy, D.C. Peterson, J. Vionnet, R. Saksena, W.F. Vann, Chemoenzymatic synthesis of conjugatable oligosialic acids, Biocatal. Biotransform. 28 (2010) 41–50. [37] W. Jing, P.L. DeAngelis, Synchronized chemoenzymatic synthesis of monodisperse hyaluronan polymers, J. Biol. Chem. 279 (2004) 42345–42349. [38] W.T. Forsee, R.T. Cartee, J. Yother, Role of the carbohydrate binding site of the Streptococcus pneumoniae capsular polysaccharide type 3 synthase in the transition from oligosaccharide to polysaccharide synthesis, J. Biol. Chem. 281 (2006) 6283–6289.

Universal fluorescent acceptor for polySTs / T.G. Keys et al. / Anal. Biochem. 427 (2012) 107–115 [39] L. Masson, B.E. Holbein, Physiology of sialic acid capsular polysaccharide synthesis in serogroup B Neisseria meningitidis, J. Bacteriol. 154 (1983) 728–736. [40] J.F. May, R.A. Splain, C. Brotschi, L.L. Kiessling, A tethering mechanism for length control in a processive carbohydrate polymerization, Proc. Natl. Acad. Sci. USA 106 (2009) 11851–11856. [41] S.P. Galuska, R. Geyer, R. Gerardy-Schahn, M. Mühlenhoff, H. Geyer, Enzymedependent variations in the polysialylation of the neural cell adhesion molecule (NCAM) in vivo, J. Biol. Chem. 283 (2008) 17–28. [42] S.P. Galuska, I. Oltmann-Norden, H. Geyer, B. Weinhold, K. Kuchelmeister, H. Hildebrandt, R. Gerardy-Schahn, R. Geyer, M. Mühlenhoff, Polysialic acid profiles of mice expressing variant allelic combinations of the polysialyltransferases ST8SiaII and ST8SiaIV, J. Biol. Chem. 281 (2006) 31605–31615. [43] P.M. Drake, J.K. Nathan, C.M. Stock, P.V. Chang, M.O. Muench, D. Nakata, J.R. Reader, P. Gip, K.P. Golden, B. Weinhold, R. Gerardy-Schahn, F.A. Troy 2nd, C.R. Bertozzi, Polysialic acid, a glycan with highly restricted expression, is found on human and murine leukocytes and modulates immune responses, J. Immunol. 181 (2008) 6850–6858. [44] H. Hildebrandt, M. Mühlenhoff, I. Oltmann-Norden, I. Röckle, H. Burkhardt, B. Weinhold, R. Gerardy-Schahn, Imbalance of neural cell adhesion molecule and polysialyltransferase alleles causes defective brain connectivity, Brain 132 (2009) 2831–2838.

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[45] K. Angata, M. Suzuki, M. Fukuda, ST8Sia II and ST8Sia IV polysialyltransferases exhibit marked differences in utilizing various acceptors containing oligosialic acid and short polysialic acid, J. Biol. Chem. 277 (2002) 36808–36817. [46] J.L. Zapater, K.J. Colley, Sequences prior to conserved catalytic motifs of the polysialyltransferase, ST8SiaIV, are required for substrate recognition, J. Biol. Chem. 287 (2012) 6441–6453. [47] D. Nakata, L. Zhang, F.A. Troy, Molecular basis for polysialylation: a novel polybasic polysialyltransferase domain (PSTD) of 32 amino acids unique to the a2,8-polysialyltransferases is essential for polysialylation, Glycoconj. J. 23 (2006) 423–436. [48] G.T. Barry, Colominic acid, a polymer of N-acetylneuraminic acid, J. Exp. Med. 107 (1958) 507–521. [49] E.J. McGuire, S.B. Binkley, The structure and chemistry of colominic acid, Biochemistry 3 (1964) 247–251. [50] A.K. Bhattacharjee, H.J. Jennings, C.P. Kenny, A. Martin, I.C. Smith, Structural determination of the sialic acid polysaccharide antigens of Neisseria meningitidis serogroups B and C with carbon 13 nuclear magnetic resonance, J. Biol. Chem. 250 (1975) 1926–1932. [51] W. Egan, T.-Y. Liu, D. Dorow, J.S. Cohen, J.D. Robbins, E.C. Gotschlich, J.B. Robbins, Structural studies on the sialic acid polysaccharide antigen of Escherichia coli strain Bos-12, Biochemistry 16 (1977) 3687–3692.